Comparative Characterization of the Major Human

COMPARATIVE CHARACTERIZATION OF THE MAJOR HUMAN

GLUTAREDOXIN ISOZYMES AND IDENTIFICATION OF A MECHANISM

BY WHICH GRX1 REGULATES APOPTOSIS IN CARDIOMYOCYTES

MOLLY MEGAN GALLOGLY

Submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy

Dissertation Advisor: Dr. John J. Mieyal

Department of Pharmacology

CASE WESTERN RESERVE UNIVERSITY

August, 2009

CASE WESTERN RESERVE UNIVERSITY

SCHOOL OF GRADUATE STUDIES

We hereby approve the thesis/dissertation of

______Molly Megan Gallogly______candidate for the ______Ph.D.______degree *.

(signed) ______Charles L. Hoppel, M.D.______

(chair of the committee)

______John J. Mieyal, Ph.D.______

______Clark W. Distelhorst, M.D.______

______George R. Dubyak, Ph.D.______

______Michael E. Maguire, Ph.D.______

______

(date) ______June 11, 2009______

*We also certify that written approval has been obtained for any proprietary material

contained therein.

Dedication

To my husband, Timothy John Miller, for experiencing every step on my way to the PhD

(both forward and back!) alongside me; for encouraging me when I was sure I would

never finish; for challenging me to try things I never would have considered, seeing

potential in me that I didn’t know I had; and for being the best husband and father

he can be, making our family the greatest gift of my graduate career.

TABLE OF CONTENTS

LIST OF SCHEMES...... 4 LIST OF TABLES ...... 5 LIST OF FIGURES ...... 6 LIST OF ABBREVIATIONS ...... 8 ACKNOWLEDGMENTS ...... 15 ABSTRACT ...... 21 CHAPTER 1: BACKGROUND AND INTRODUCTION ...... 23 1.1 INTRODUCTION TO REVERSIBLE PROTEIN GLUTATHIONYLATION 24 1.1.1 Summary of Recent Advances ...... 24 1.1.2 Introduction ...... 25 1.1.3 Chemistry of Protein S-Glutathionylation and Deglutathionylation ...... 26 1.1.4 Catalysis of protein deglutathionylation ...... 32 1.1.5. Catalysis of Protein Glutathionylation ...... 37 1.1.6 Other potential mechanisms of catalysis of protein S-glutathionylation ...... 40 1.1.7 Conclusions & Frontiers ...... 41 1.2 MECHANISTIC DETAILS OF CATALYSIS OF THIOL-DISULFIDE EXCHANGE BY GLUTAREDOXIN ...... 43 1.2.1 Introduction ...... 43 1.2.2 Overall catalytic scheme for glutaredoxin ...... 43 1.2.3. Characteristics of individual steps of glutaredoxin catalysis of thiol-disulfide exchange ...... 45 1.2.4 Evaluation of the mechanisms of the catalytic enhancement by glutaredoxin 49 1.2.5 Reports of complex formation between Grx and GSH ...... 52 1.2.6 Catalysis of Deglutathionylation by Other Glutaredoxins and Glutaredoxin Domains ...... 55 1.3 REGULATION OF GLUTAREDOXIN ACTTVITY ...... 65 1.3.1 Subcellular Localization and Effects on Grx Activity ...... 65 1.3.2. Enzyme concentration ...... 67 1.3.3. Chemical environment/milieu ...... 68 1.3.4 Temporal Regulation of Glutaredoxin Activity...... 70

1.3.5 Conclusion ...... 80 1.4 GLUTATHIONYLATION AND GLUTAREDOXINS IN CARDIOVASCULAR DISEASE ...... 81 1.4.1 Cardiovascular Diseases and Alterations in Protein-S-glutathionylation Status ...... 81 1.4.2 Myocardial Infarction ...... 82 1.4.3 Preconditioning ...... 90 1.4.4 Nonspecific oxidative injury ...... 95 1.4.5 Cardiac Hypertrophy ...... 99 1.4.6. Atherosclerosis ...... 100 1.5 REGULATION OF APOPTOSIS BY GLUTATHIONYLATION AND GLUTAREDOXINS ...... 107 1.6 BCL-2 AND BCL-XL ...... 108 CHAPTER 2: HYPOTHESES, AIMS, EXPERIMENTAL APPROACHES, AND OVERVIEW OF FINDINGS ...... 137 2.1. KINETIC CHARACTERIZATION OF MAMMALIAN GRX2 ...... 137 2.2. REGULATION OF APOPTOSIS IN CARDIOMYOCYTES BY GRX1 VIA NFΚB AND TARGET GENES BCL-2 AND BCL-XL ...... 142 2.2.1 JUSTIFICATION OF THE MODEL SYSTEMS ...... 146 CHAPTER 3: KINETIC AND MECHANISTIC CHARACTERIZATION AND VERSATILE CATALYTIC PROPERTIES OF MAMMALIAN GLUTAREDOXIN 2: IMPLICATIONS FOR INTRACELLULAR ROLES ...... 150 3.1 Abstract ...... 150 3.2 Introduction ...... 151 3.3 Materials and Methods ...... 154 3.3.1 Materials ...... 154 3.3.2 General Methods...... 157 3.3.3 Specific Methods ...... 157 3.4 Results ...... 166 3.5 Discussion ...... 174 CHAPTER 4: GLUTAREDOXIN (GRX1) REGULATES APOPOTOSIS IN CARDIOMYOCYTES VIA NFΚB TARGETS BCL-2 AND BCL-XL: IMPLICATIONS FOR CARDIAC AGING ...... 208

4.1 Abstract ...... 208 4.2 Introduction ...... 209 4.2. Materials and Methods ...... 211 4.2.1. General materials ...... 211 4.2.2. Animals ...... 212 4.2.3. Specific Methods ...... 212 4.3. Results ...... 223 4.4 Discussion ...... 228 4.5 Conclusion ...... 233 CHAPTER 5: CONCLUSIONS AND FUTURE DIRECTIONS ...... 252 5.1 KINETIC CHARACTERIZATION AND POTENTIAL PHYSIOLOGICAL ROLES OF GRX2 ...... 252 5.1.1 Conclusions and Remaining Questions ...... 252 5.1.2 Is Grx2 active as a deglutathionylating enzyme in situ in resting cells? How much does it contribute to total cellular deglutathionylation activity? ...... 254 5.1.3 What regulates the release of Grx2 from 2Fe2S cluster dimers? Does dimer dissociation correspond to increased deglutathionylation activity as expected? ..... 255 5.1.4 What are the physiological substrates for Grx2? ...... 259 5.1.5 What could be the physiological role(s) of Grx2-2Fe2S in mitochondria? ... 262 5.2 Regulation of cardiomyocyte apoptosis by Grx1 via modulation of NFκB- dependent transcription of Bcl-2 and Blc-xL ...... 264 5.2.1 Conclusions and Remaining Questions ...... 264 5.2.2 What is the site of regulation of the NFκB pathway by Grx1? ...... 265 5.2.3 Determine relative levels of Bcl-2 and Bcl-xL in adult vs. elderly F344 rats 271 5.2.4 Can elderly cardiomyocytes be “rescued” from their apoptotic phenotype by addition of Grx1 or Bcl-xL? ...... 274 5.2.5 Potential role for Grx1 and/or Bcl-xL as therapeutic targets in the aging heart ...... 279 BIBLIOGRAPHY ...... 283

LIST OF SCHEMES

Scheme Title page

1.1 Potential spontaneous chemical reactions leading to protein 109 glutathionylation 1.2 Reactions of mammalian glutaredoxins 110 1.3 Proposed mechanism for Prx glutathionylation by GSTπ 113 1.4 Complete commitment to catalysis vs. encounter-type catalytic 114 mechanisms 2.1 Model of Grx1-mediated regulation of apoptosis in cardiomyocytes 148 3.1 Catalytic mechanisms of mammalian glutaredoxin (Grx) and 183 thioredoxin (Trx)

LIST OF TABLES

Table Title page

1.1 Candidate enzymes for catalysis of protein S-glutathionylation 115 1.2 Deglutathionylation activities and glutathionyl specificity of 117 glutaredoxins from prototype organisms 1.3 Potential phosphorylation sites of human Grx enzymes 120 1.4 Cardiovascular proteins regulated by S-glutathionylation 122 1.5 Evidence for regulation of apoptosis by Grx isoforms 124 1.6 Proposed functions of Bcl-2 and Bcl-xL proteins 126 3.1 S-methylglutathione does not inhibit Grx2-catalyzed 185 deglutathionylation 3.2 Catalytic efficiencies of Grx isoforms 186 3.3 Specific activities of Grx isoforms for model sybstrate cysteine-SSG 188 (CSSG) 3.4 Helix 2 dipole moments for hGrx1 and hGrx2 189 3.5 Enhancement of GSH nucleophilicity by Grx isoforms 190 3.6 Contribution of TR to Grx-mediated deglutathionylation of CSSG 192 3.7 hGrx1 and hGrx2 exhibit equal rates of protein glutathionylation in the 193 presence of GS and GSNO, but not GSSG 5.1 Apoptotic regulators exhibiting substantial changes in mRNA content 282 with Grx1 knock-down

LIST OF FIGURES

Figure Title page

1.1 Inactivation of PTP1B by QSOX/GSH 127 1.2 Grx residues reported to interact with a covalently bound GS-moiety 128 in the Grx-SSG mixed disulfide 1.3 Predicted phosphorylation sites for human Grx isoforms 130 1.4 Proposed interactions between Grx1 and cytosolic enzymes 132 1.5 Examples of acute regulation of Grx1 133

1.6 Downstream effects of Ras regulation in response to endogenous H2O2 134 production 1.7 Downstream effects of Ras glutathionylation in response to exogenous 136 peroxynitrite or oxLDL 2.1 Sites of regulation of the NFκB pathway by reversible 149 glutathionylation 3.1 Characterization of recombinant mammalian Grx2 194 3.2 hGrx1 and hGrx2 are selective for protein-GSH mixed disulfide 196 substrates 3.3 Two-substrate kinetic analysis of hGrx2- and mGrx2-catalyzed 198 deglutathionylation demonstrates a ping-pong kinetic pattern

3.4 hGrx2 (C40S) exhibits higher KM and Vmax for CSSG, but lower KM 202 for GSH, compared to wild type enzyme 3.5 pH rate profile of BSA-SSG deglutathionylation in the absence and 204 presence of hGrx2

3.6 Determination of the pKa of the active site Cys-SH of hGrx2 205 3.7 Thiyl radical (GS•) scavenging activity of Grx2 206 4.1 Grx1 content and activity in heart tissue cytosol and in isolated 235 myocytes from F344 rats

Figure Title page

4.2 Grx1 content and total Grx activity are decreased via transfection of targeted siRNA into H9c2 cells 237 4.3 Grx1 knock-down increases apoptotic susceptibility in H9c2 cells 239 4.4 Grx1-deficient cells exhibit decreased NFκB transcriptional activity 241 without significant change in NFκB content 4.5 Effect of IKK inhibitor BMS 345541 on NFκB activity and apoptotic 243 susceptibility in wild-type H9c2 cells 4.6 Normalized amounts of Bcl-2 and Bcl-xL mRNA in Grx1 knock-down 245 cells relative to control cells 4.7 Contents of Bcl-2 and Bcl-xL proteins are decreased in Grx1-deficient 246 H9c2 cells. 4.8 Effect of Bcl-2 and Bcl-xL knock-down on apoptotic susceptibility in 248 H9c2 cells. 4.9 NFκB-dependent transcriptional activity in cardiomyocytes isolated 251 from adult and elderly F344 rat hearts

LIST OF ABBREVIATIONS

2Fe2S 2-iron, 2-sulfur cluster

6-mo 6 month-old

24-mo 24 month-old

αKGDH α-ketoglutarate dehydrogenase

βME β-mercaptoethanol

βME-SSG β-mercaptoethanol-glutathione mixed disulfide

AII angiotensin II

Ab antibody

ADP adenosine diphosphate

Akt protein kinase B

AMPSO N-(1,1-dimethyl-2-hydroxyethyl)-3-amino-2-hydroxypropanesulfonic

acid

ANT adenine nucleotide translocase

AP-1 activator protein-1 apoB100 apolipoprotein B100

ARE antioxidant response element

ASK1 apoptosis signal regulating kinase 1

ASO arteriosclerosis obliterans

BAEC bovine aortic endothelial cells

Bcl-2 B cell CLL/lymphoma 2

Bcl-xL B cell lymphoma x long

BCNU bis-chloronitrosourea

BMS 345541 (4(2-aminoethyl)amino-1,8-dimethylimidazo(1,2-a)quinoxaline)

BSA bovine serum albumin

BSA-SSC bovine serum albumin-cysteine mixed disulfide

BSA-SSG bovine serum albumin-glutathione mixed disulfide

C40S cysteine 40 to serine mutant

CA-III carbonic anhydrase 3

CARD10 caspase recruitment domain family member 10

Casp8ap2 caspase 8-associated protein 2

CDDO-Im 2-cyano-3,12 dioxooleana-1,9 dien-28-imidazolide

CHX cycloheximide

CSSG cysteine-glutathione disulfide

Cys cysteine

DAPI 4',6-diamidino-2-phenylindole

Dffb DNA fragmentation factor beta subunit

DMEM Dulbecco’s modified Eagle medium

DNA deoxyribonucleic acid

DTNB 5,5′-dithiobis-(2-nitrobenzoic acid)

DTT dithiothreitol

EDTA ethylenediaminetetraacetic acid

EGF epidermal growth factor eNOS endothelial nitric oxide synthase

Ec Escherichia coli

EPR electron paramagnetic resonance

ER endoplasmic reticulum

EST expressed sequence tag

F344 Fischer 344

FAD flavin adenine dinucleotide

FBS fetal bovine serum

FGF fibroblast growth factor

Gadd45 growth arrest and DNA damage inducible 45 alpha

GAPDH glyceraldehyde phosphate dehydrogenase

GATA4 GATA-binding protein 4

GFP green fluorescent protein

GR glutathione disulfide reductase

Grx glutaredoxin

GS- glutathionyl moiety

GS• glutathione thiyl radical

GSNO S-nitrosoglutathione

GS(O)SG glutathione thiosulfinate

GSSG glutathione disulfide h human

HEDS hydroxyethyl disulfide

HEK human embryonic kidney

HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid

HEPPSO N-[2-hydroxyethyl]piperazine-N′-[2-hydroxypropanesulfonic acid]

HPLC high performance liquid chromatography

HRP horseradish peroxidase

IκB inhibitor of κB

IAM iodoacetamide

IKK inhibitor of κB (IκB) kinase

IL interleukin

IP immunoprecipitation

IP3R inositol 1,4,5-triphosphate receptor

IPC ischemic preconditioning

IR ischemia-reperfusion

IRS insulin receptor substrate

JNK Jun N-terminal kinase

KD knock-down kDa kilodalton

KM Michaelis constant; concentration of substrate at which v = ½ Vmax

int KM intrinsic (or “true”) KM

Kmix The specific oxidation potential (Kox) for formation of the mixed disulfide of protein and glutathione (protein-SSG)

KO knockout

Kox the GSH:GSSG ratio at which protein-SH:protein-SSG = 1 m mouse

Mapk8ip mitogen-activated protein kinase 8 interacting protein

MES 2-(N-morpholino)ethanesulfonic acid

MI myocardial infarction

MMP matrix metalloproteinase precursor

MOI multiplicity of infection

MPTP 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine

NADPH nicotinamide adenine dinucleotide phosphate

NEM N-ethylmaleimide

NFκB nuclear factor κB

NMR nuclear magnetic resonance

NP-40 nonyl phenoxylpolyethoxylethanol

NTSP N-succinimidyl pyridyl bis-(3,3’-dithiopropionate)

- O2 superoxide

OH hydroxyl radical

ONOO- peroxynitrite oxLDL oxidized LDL

PABA/NO O2-[2,4-dinitro-5-(N-methyl-N-4-carboxyphenylamino)phenyl] 1-N,N-

dimethylamino)diazen-1-ium-1,2-diolate

PAGE polyacrylamide gel elecrophoresis

PC proprotein convertase

PICOT protein kinase C theta (θ)-interacting cousin of thioredoxin

PKC protein kinase C

Prx peroxiredoxin

Protein-SSG protein-GSH mixed disulfide

PSG protein with a reduced cysteine sulfhydryl moiety

PSNO S-nitrosylated protein

PTEN phosphatase and tensin homolog

PTP1B protein tyrosine phosphatase 1B

PVDF polyvinylidene difluoride

QSOX quiescein sulfhydryl oxidase

RLU relative light units

RNS reactive nitrogen species

RT-PCR real-time PCR (polymerase chain reaction)

ROS reactive oxygen species

RyR ryanodine receptor

Sc Saccharomyces cerevisiae

SDS sodium dodecyl sulfate

SEM standard error of the mean

SERCA sarcoplasmic/endoplasmic reticulum calcium ATPase

SIN-1 3-morpholinosydnonimine sIR simulated ischemia-reperfusion shRNA small hairpin RNA siRNA small interfering RNA

SOD superoxide dismutase

SPDP N-Succinimidyl 3-(2-pyridyldithio)-propionate

SQR succinate ubiquinone reductase

SRx sulfiredoxin

TCA trichloroacetic acid

Tdh3 S. cerevisiae homolog of GAPDH

TDOR thiol-disulfide oxidoreductase

TG transgenic

TNF tumor necrosis factor

TR thioredoxin reductase

Trp transformation-related protein

Trx thioredoxin

Vmax maximal reaction rate (velocity)

int Vmax intrinsic (“true”) Vmax

VSMC vascular smooth muscle cells

WT wild-type

X xanthine

XO xanthine oxidase

ACKNOWLEDGMENTS

I am deeply grateful for valuable scientific and personal support I received from many people during my time as a graduate student in the Department of Pharmacology.

Dr. John Mieyal, my thesis advisor, has served as an exemplary model of careful and rigorous scientific inquiry, always striving to examine data objectively and to base experimental decisions on reproducible observations. I appreciate Dr. Mieyal’s dedication to making time for his students, always ensuring an hour a week for one-on- one discussions, regardless of administrative obligations or grant deadlines. He has always expressed interest in my life outside of the laboratory, asking about my family and remaining sensitive to personal concerns. Outside his office, Dr. Mieyal has served as an effective mediator, exhibiting an enviable ability to remain calm and constructive in the midst of conflict. He shows his joy for life by seeking new and exciting experiences, and inspires me to see the best in the people around me. To be sure, working under his direction has benefitted me both scientifically and personally.

Other members of the Mieyal laboratory have helped tremendously in the development of my thesis work. David Starke taught me to become a kineticist by discussing enzymology theory, reviewing data with painstaking attention to detail, and assisting with the design of critical experiments. I greatly enjoyed the scientific discourse we shared, and appreciate the education I received in protein purification, enzyme behavior, and every once in a while, the great game of baseball. Sue Ospina performed the initial comparison of specific activity between hGrx1 and hGrx2, as well as preliminary kinetic analysis of human and mouse Grx2. She also provided key tips on

the purification of glutaredoxin, and offered information and assistance freely and kindly.

Amanda Leonberg, a former research assistant, performed 2-substrate kinetics and pKa

analysis on hGrx2, and was a kind and considerate coworker for the short time she was a

part of our laboratory. Dr. Harish Pai, a former post-doc, spearheaded efforts to knock down Grx1 stably in H9c2 cells, and determined the apoptotic susceptibility of the knock- down cells to oxidative challenges. He also conducted pilot studies on NFκB activity in

H9c2 cells with stable Grx1 knock-down, and on the effect of the IKK inhibitor Bay 11-

7085 on apoptotic susceptibility in wild-type cells. Shortly after her arrival in the

laboratory, Mary Consolo took on purification of hGrx2, which facilitated completion of

kinetic studies of the enzyme. Mary also made everyone’s life in the lab a little more

bearable each day, keeping us laughing during the most stressful times. Dr. Suparna

Qanungo taught me how to use hypoxic chambers, and performed Western blot analysis

for Trx on heart tissue from adult and elderly rats. Suparna has also proven a steadfast

friend and supporter since joining the laboratory, offering the welcome perspective of

another mom in science seeking the perfect balance between work and family. Dr.

Melissa Shelton determined NFκB activity in Grx1 knock-down cells, and developed the

protocol for determination of NFκB activity in primary cardiomyocytes. Melissa also

helped me to design my first siRNA knock-down experiments in H9c2 cells. Sarah

Stewart performed isolations of primary cardiomyoctes from adult and elderly Fischer

344 rats, and Elizabeth Sabens took pictures of these cardiomyocytes shown in Chapter 4.

Several members of the Case Comprehensive Cancer Center facilitated work described in Chapter 4. Jose Gomez and Dr. Vivian Gama (Dr. Shigemi Matsuyama’s laboratory, Department of Pharmacology) taught me to do Hoechst staining by providing

protocols and reagents and generously donating their time to demonstrate the technique

on my cells. Dr. Michael Davis (Dr. Clark Distelhorst’s laboratory, Department of

Pharmacology) taught me how to measure calcium levels in H9c2 cells, and helped me

perform sub-G1 flow cytometry in Case’s core facility. Although these experiments are

not described here, performing them introduced me to unfamiliar techniques and taught

me how to approach scientific questions relating to calcium homeostasis and cell death.

Other members of the Distelhorst Laboratory provided personal support, reagents, and a

valuable “open-door” policy during my graduate school career. Karen McColl offered

protocols, antibodies, and equipment on-demand. Jason Molitoris helped me perform

RT-PCR for Bcl-2 in H9c2 cells, and continued to lend me RNeasy kits as I pursued PCR array analysis of mRNA from these cells. Dr. Distelhorst himself freely offered the time and skills of his laboratory personnel, and his generosity allowed me to explore areas of science I never would have otherwise considered. Finally, Vaibhav Patakh (CWRU Gene

Expression & Genotyping Core Facility; Dr. Martina Veigl, Director) performed PCR array analysis on mRNA from control and Grx1 knock-down H9c2 cells. Vai’s friendly and helpful attitude, combined with his conscientious approach to my experiment, facilitated completion of an experiment unfamiliar to our laboratory.

I was fortunate to receive help from scientists outside of Case Western Reserve

University with certain aspects of my thesis work. Drs. Periannan Kuppusamy and

Mahmood Khan (Dorothy M. Davis Heart and Lung Research Institute, The Ohio State

University, Columbus, OH) assisted Dr. Suparna Qanungo in the detection of glutathionyl radical (GS) by EPR spectroscopy. These studies confirmed GS formation

via the radical-generating system described in Chapter 3. Drs. David Roth and Hemal

Patel (University of California, San Diego, Department of Anesthesiology) shared their protocol for isolation and culture of primary cardiomyocytes, which served as the basis for our optimized procedure (see Chapter 4) and helped members of our laboratory to optimize our protocol. Dr. Edward Lesnefsky (formerly of the Louis Stokes Veterans

Affairs Medical Center, Cleveland, OH; now of McGuire Veterans Affairs Medical

Center, Richmond, VA) greatly facilitated development of our cardiomyocyte isolation protocol by initiating the collaboration with Drs. Roth and Patel, and by making his laboratory available during all stages of methods development.

My thesis committee (Dr. Charles Hoppel (chair), Dr. Michael Maguire, Dr. George

Dubyak, Dr. Clark Distelhorst, Dr. John Mieyal) has provided sound and practical feedback on both of my projects over the last five years. Dr. Hoppel’s insights on the aging heart project, as well as my transition back to the clinic, are most appreciated. I am also grateful for helpful conversations with Drs. Dubyak and Distelhorst focused on development of experimental strategies and relevance of recent publications in the fields of redox regulation, calcium homeostasis, and Bcl-2 family proteins.

I thank the Medical Scientist Training Program for supporting so many years of my medical and graduate education. I appreciate that Directors Dr. Cliff Harding and Dr.

George Dubyak always made themselves available to discuss my questions and concerns.

I also thank the administrative staff (Kathy Schultz, Donna McIlwain, Deidre Gruning, and Bart Jarmusch) for helping to make an incredibly rich and supportive program run smoothly; for welcoming questions and never hesitating to solve a student’s problem.

Most of all, I thank the women of the MSTP (especially Sarah Drawz and Bridgette

Christopher) for being great friends, inspiring role models, and the most incredible

support system I could have asked for. These amazing women have helped me navigate

scientific and personal struggles so that I could realize my strengths and recognize my

accomplishments in the midst of seemingly insurmountable obstacles. Other friends in

Case’s medical and graduate programs have provided unconditional and much-

appreciated support, especially Sarah Swerdlow and Dr. Victor Ibrahim.

I have been fortunate to receive valuable guidance from other mentors involved in the

clinical and basic sciences. Dr. Kevin Evans, my undergraduate chemistry advisor, was

the first to suggest I consider scientific research. Had he not offered me a position on his

research team, I might never have considered pursuing a PhD. I am grateful not only for

his interest in my academic development, but also my personal growth and happiness

both at Grinnell College and at Case Western Reserve University. My clinical mentors,

Dr. Marjorie Greenfield and Dr. Reshma Shah, welcomed me into their practices and

offered valuable perspectives on their respective clinical fields. I am grateful to call Dr.

Greenfield both my friend and a great clinical inspiration, introducing me to the life of

someone “on the other side,” and serving as a great example of a patient-friendly, conscientious physician who has achieved an admirable work-life balance. Other

“unofficial” mentors who have offered me valuable advice include Dr. Robert Haynie, my society dean, and Dr. Colleen Croniger (CWRU Dept. of Nutrition).

My family’s support has been critical and unfailing since I entered graduate school.

My parents, Jill and Tim Gallogly, gave me the gift of their full confidence, always wanting what was best for me, and supporting whatever path I chose to figure out what that is. I thank my sisters (biological sister Sarah Gallogly, sister-in-law Karen Miller

Song, and brain-twin sister Rachel Melis) for being open ears during some of my hardest

19 trials. I am grateful to my new relatives, the Millers and Wiedemers, for welcoming me into their lives and sharing the joy of our growing family. Most of all, I thank my husband, Tim Miller, and son Gavin Bryce Miller, for showing me the joys of life outside of the lab, and for always encouraging me to stop, drop and laugh.

Comparative Characterization of the Major Human

Glutaredoxin Isozymes and Identification of a Mechanism

by which Grx1 Regulates Apoptosis in Cardiomyocytes

Abstract

MOLLY MEGAN GALLOGLY

Mixed disulfides between cysteines and glutathione (GSH) (i.e., protein S- glutathionylation) represent an oxidative post-translational modification with roles in sulfhydryl homeostasis and signal transduction. Glutaredoxin (Grx) is an efficient

catalyst of protein deglutathionylation, and human Grx1 (cytosolic isoform) is the

prototype enzyme. Recently, a second isoform (Grx2) was discovered. Systematic

analysis of the human glutaredoxins demonstrated remarkable similarities, including

specificity for glutathionyl mixed disulfides and double displacement, encounter-type

catalysis where reduction of the Grx-SSG intermediate is rate-determining. Both Grx1

and Grx2 are recycled preferentially by GSH, and both promote protein glutathionylation

with activated donors GS and GSNO. However, Grx2 has 10-fold lower catalytic

efficiency than Grx1, due to decreased Vmax. Lower rate enhancement by Grx2 is explained by (a) 1 pH unit increase in pKa of the catalytic cysteine serving as leaving

group in the rate-determining step; and (b) less enhancement of GSH nucleophilicity

towards the Grx-SSG intermediate. Calculated concentrations of Grx1 and Grx2 suggest

equal deglutathionylase activities for the subcompartments of mitochondria, and

potentially more GS-mediated glutathionylation by Grx2 in the matrix. Redox

regulation of apoptosis is an area of intense study. We examined Grx1 regulation of

apoptosis in cardiomyocytes, which exhibit increased apoptotic susceptibility and

diminished Grx1 with aging in animal models. To determine whether Grx1 deficiency

increases apoptotic susceptibility, Grx1 was knocked down in H9c2 cardiomyocytes to a

similar extent as in cardiomyocytes from elderly animals. Grx1 knock-down resulted in

increased apoptosis at baseline and following oxidative stress; concomitantly NFκB

transcriptional activity was diminished, and its anti-apoptotic gene products Bcl-2 and

Bcl-xL were decreased. Inhibiting NFκB, or knocking down Bcl-2 and/or Bcl-xL in

wild-type H9c2 cells to the same extent as in Grx1 knock-down cells increased apoptosis

at baseline; but only Bcl-xL knock-down increased oxidant-induced apoptosis and fully

accounted for the apoptotic phenotype observed in Grx1-deficient cells. NFκB activity

was diminished also in cardiomyocytes from elderly Fischer 344 rats, indicating

correspondence of the cellular and animal models of aging. This thesis provides

foundations for future investigations on: (a) specific roles of glutaredoxins in sulfhydryl

regulation within mitochondria; and (b) mechanisms of Grx1-dependent regulation of apoptotic signaling in cardiomyocytes.

CHAPTER 1: BACKGROUND AND INTRODUCTION

Portions of this chapter are reproduced from Current Opinion in Pharmacology 2007 7(4):381-91, Copyright 2007; Antioxidants & Redox Signaling 10(11):1941-88, Copyright 2008; and Antioxidants & Redox Signaling 11(5):1059-1081, Copyright 2009. Reproduced with permission from Elsevier Publishers and Mary Ann Liebert Publications.

Protein glutathionylation is an oxidative post-translational modification consisting of

disulfide formation between the sulfhydryl group of a protein cysteine and the sulfhydryl

of the small intracellular tripeptide, glutathione (GSH, i.e., γ-glutamylcysteinylglycine).

Glutathionylation serves two critical roles in cells: protection of cysteine residues from irreversible oxidation, and redox signal transduction. In many cases, glutathionylation alters protein function; thus, reversible glutathionylation of specific proteins has been implicated in the regulation of proteins involved in metabolism, cytoskeletal reorganization, cellular hypertrophy, inflammation, and apoptosis (Mieyal et al., 2008).

Glutaredoxin (Grx) is a thiol-disulfide oxidoreductase (TDOR) enzyme that specifically

and efficiently catalyzes GSH-dependent reduction of glutathionylated proteins (protein-

SSGs), and mammalian Grx regulates diverse cellular functions via deglutathionylation

of target protein-SSGs (Mieyal et al., 2008). Under certain conditions, mammalian

glutaredoxins can facilitate protein glutathionylation, and the recently characterized Grx2

isoform may play a role also in iron-sulfur cluster biosynthesis (Gallogly et al., 2009).

To date, deglutathionylation remains the best characterized catalytic function of Grx enzymes, and evidence for the physiological significance of Grx-catalyzed deglutathionylation in human disease continues to grow (Mieyal et al., 2008; Dalle-

Donne et al., 2007).

This chapter sets the stage for the description of kinetic and mechanistic studies of

human glutaredoxin isoforms in Chapters 3-4 by presenting the chemical mechanisms for

reversible protein glutathionylation (Chapter 1.1), details of catalysis of thiol-disulfide exchange by glutaredoxins (Chapter 1.2), and potential mechanisms for regulation of Grx activity (Chapter 1.3). Discussions of the significance of Grx in cardiovascular disease

(Chapter 1.4), and in the regulation of apoptosis (Chapter 1.5) via Bcl-2 and Bcl-xL

(Chapter 1.6), are particularly pertinent to the mechanistic studies of apoptotic regulation by Grx1 in cardiomyocytes described in Chapter 4.

1.1 INTRODUCTION TO REVERSIBLE PROTEIN GLUTATHIONYLATION

1.1.1 Summary of Recent Advances

Reversible protein S-glutathionylation (protein-SSG) is an important posttranslational modification in two contexts: it protects protein cysteines from irreversible oxidation and it represents an important regulatory mechanism in redox signal transduction. Analogous to the dephosphorylation role of phosphatases in modulating signal transduction, enzymes of the glutaredoxin (Grx) family of thiol-disulfide oxidoreductases catalyze deglutathionylation of various specific protein substrates, exerting regulatory effects on diverse intracellular signaling pathways (Finkel, 2000; Klatt and Lamas, 2000; Shelton et al., 2005). Recently, other enzymes have been reported to exhibit deglutathionylating activity; however, the extent of their contribution to intracellular protein deglutathionylation is uncertain. Currently, no enzyme has been shown to serve as a catalyst of S-glutathionylation in situ, although potential prototypes are reported, including human Grx1 and the π isoform of glutathione-S-transferase (GSTπ). Further insight on cellular mechanisms of protein glutathionylation and deglutathionylation will

enrich our understanding of redox signal transduction pathways and potentially identify

new therapeutic targets for diseases in which oxidative stress perturbs normal redox

signaling. Accordingly, this overview focuses primarily on mechanisms of catalysis in

mammalian systems.

1.1.2 Introduction

S-glutathionylation of proteins is a reversible post-translational modification with

roles in sulfhydryl homeostasis and signal transduction. Protein glutathionylation

increases globally during overt oxidative stress (e.g., cardiac ischemia-reperfusion,

(Eaton et al., 2002)), but selectively in the presence of reactive oxygen species (ROS) that are generated during physiologic signaling (e.g., growth factor or angiotensin II receptor occupation (Wang et al., 2001b; Wingert et al., 2005; Adachi et al., 2004a)).

Reversible glutathionylation modulates the activities of diverse protein substrates in situ,

with effects on multiple physiological events, including cell proliferation (Kanda et al.,

2006), cytoskeletal organization (Wang et al., 2003), protein synthesis (Adachi et al.,

2004a), transcription (Shelton et al., 2007; Reynaert et al., 2006), and vascular tension

(Adachi et al., 2004b).

As a guide for evaluating reports of potential regulation via reversible glutathionylation, we have proposed 5 criteria for establishing protein glutathionylation as a regulatory mechanism (Shelton et al., 2005). Briefly stated, S-glutathionylation must

(1) change the function of the modified protein, (2) occur in intact cells in response to a physiological stimulus and lead to a physiological response, (3) occur at relatively high

GSH:GSSG ratios (i.e., physiological conditions), and exhibit rapid and efficient mechanisms for (4) formation and (5) reversal of protein-SSG. To date studies of three

proteins stand out as best fulfilling these criteria for regulation, namely, actin (Wang et

al., 2001b; Wang et al., 2003), Ras (Adachi et al., 2004a), and the protein tyrosine

phosphatases (PTPs) (Kanda et al., 2006; Barrett et al., 1999a; Barrett et al., 1999b), but

the mechanism(s) of their glutathionylation in vivo remains unclear (i.e., criterion (#4)).

Protein deglutathionylation is efficiently catalyzed by the thiol-disulfide oxidoreductase glutaredoxin (Grx), and it has been shown to account for most of the the deglutathionylating activity in mammalian cells (Chrestensen et al., 2000; Jung and

Thomas, 1996). Potential contributions of other enzymes to physiological deglutathionylation are discussed in Section 1.1.4. Potential enzymatic mechanisms of glutathionylation are discussed in Section 1.1.5.

1.1.3 Chemistry of Protein S-Glutathionylation and Deglutathionylation

1.1.3A. Thiol-disulfide exchange

Protein glutathionylation may proceed spontaneously either by thiol-disulfide exchange (Scheme 1.1, p. 109, upper) or by reaction of a reduced protein-SH or a glutathionyl-sulfhydryl moiety with an oxidized sulfhydryl derivative (e.g., sulfenic acid, thiyl radical, etc.) (Scheme 1.1, p. 109, lower). For most protein cysteines with typical redox potentials (i.e., Kmix ~1), the intracellular GSH:GSSG ratio would have to decline

dramatically (i.e., from 100:1 to 1:1) to achieve 50% conversion of protein-SH to protein-

SSG (Gilbert, 1990). Such low GSH:GSSG ratios are difficult to achieve even under artificial oxidative stress (Gilbert, 1990; Starke et al., 1997). Therefore, although thiol- disulfide exchange can lead to protein-SSG under extreme conditions (i.e., high GSSG concentration) (Wang et al., 2001b; Barrett et al., 1999b; Pineda-Molina et al., 2001;

Beer et al., 2004), it is not a likely mechanism in vivo, because even enzyme catalysis

would not shift the position of the unfavorable equilibrium. This realization is accurately

portrayed in several previous reviews (Gilbert, 1990; Ghezzi, 2005b; Ghezzi, 2005a;

Ziegler, 1985), however it was misrepresented in a recent minireview (Sykes et al.,

2007). Importantly, there is at least one exception to this generalization, namely c-Jun,

whose unusual redox potential (Klatt et al., 1999a) makes it susceptible to modification by GSSG at relatively high GSH:GSSG ratios (i.e., 50% protein-SSG at GSH:GSSG =

13).

1.1.3B. Enhanced Protein-S-Glutathionylation via Activated Thiol-Derivatives

Likely mechanisms of protein glutathionylation within cells involve reaction of the

thiol moiety of protein-SH or GSH with a corresponding oxidized derivative, e.g., S-

nitrosyl (-SNO), sulfenic acid (-SOH), thiyl radical (-S), thiosulfinate (-S(O)SR), or

sulfenyl-amide (cyclic-S-N-CO-) (Ghezzi, 2005b; Giustarini et al., 2004; Huang and

Huang, 2002; Hurd et al., 2005; Thomas et al., 1995; Klatt and Lamas, 2000; Shelton et al., 2005) (Scheme 1.1, p. 109, lower). Production of such oxidized sulfhydryl derivatives has been suggested to occur during normal redox signaling, pathological oxidative stress, and/or treatment with xenobiotics or pharmacological agents (Di Stefano et al., 2006; Han et al., 2005; Rossi et al., 2006; Saavedra et al., 1997); exposure of

proteins and cultured cells to many of these reactive sulfhydryl intermediates leads to

protein glutathionylation in vitro (Adachi et al., 2004b; Klatt et al., 1999b; Li et al.,

2001; Mohr et al., 1999). Determining the importance of these glutathionylation

mechanisms in vivo requires evaluation of the chemistry and biochemistry, including

consideration of thermodynamic and kinetic competence, particularly under conditions

that mimic the intracellular milieu.

Sulfenic Acid Intermediates: Protein-sulfenic and glutathione-sulfenic acids are proposed to form from reaction of cysteine with endogenously produced ROS and/or reactive nitrogen species (RNS) (Barrett et al., 1999a; Huang and Huang, 2002; Poole et al., 2004). In general, sulfenic acids are considered to be highly unstable, rapidly undergoing oxidation to sulfinic (-SO2H) or sulfonic (-SO3H) acids, or reacting with

nearby thiols to form disulfides (Scheme 1.1, p. 109, lower) (Claiborne et al., 1993;

Claiborne et al., 1999), although some enzyme active sites contain uniquely stabilized

cysteine-sulfenic acids (Cys-SOH) or selenocysteine-selenenic acids (Cys-Se-OH); for

example., peroxiredoxin (PRx), glutathione peroxidase (GPx), NADH peroxidase (Npx),

methionine sulfoxide reductase (Msr)) (Poole et al., 2004; Flohe, 1985; Rhee et al.,

2005).

In the context of redox regulation, many proteins and transcription factors have been

identified as candidates for regulation by sulfenic acid formation (e.g., c-Jun, Fos, bovine papillomavirus E2 protein, nuclear factor 1, NFκB-p50, GAPDH) (Claiborne et al., 1993;

Pineda-Molina et al., 2001; Yoshitake et al., 1994). However, in most cases, the sulfenic acid was not demonstrated directly, or was observed only under artificial oxidative conditions in the absence of GSH, which would react quickly with most protein-SOH to protein-SSG (Klatt and Lamas, 2000; Poole et al., 2004). Such conversions to protein-

SSG by GSH is documented for mammalian protein tyrosine phosphatases (PTPs)

(Barrett et al., 1999a; Barrett et al., 1999b), previously reported to react with H2O2 to

form PTP1B-SOH in the absence of GSH (Lee et al., 1998, Denu and Tanner, 1998). In

fact, treatment of A431 cells with EGF (shown separately to elicit H2O2 production

intracellularly) led to glutathionylation of PTP1B in situ, specifically characterized as

C215-SSG by mass spectrometry (Barrett et al., 1999b). Thus, although protein-SOH

may be the initial form of oxidative modification for many proteins, we expect in most

cases that protein-SSG serves as the more stable intermediate in redox signaling.

S-nitrosylated intermediates: Cysteine sulfhydryls on proteins and GSH undergo

nitrosylation in vivo under a variety of normal and pathological conditions (Burwell et

al., 2006; Eiserich et al., 1998), forming protein-SNO, and GSNO, respectively. Both

GSNO and protein-SNO are relatively stable, with half lives on the order of hours in

aqueous solution in the absence of thiols (Arnelle and Stamler, 1995; Park, 1988; Singh

et al., 1996). GSNO has been detected in micromolar concentrations in non-stressed

tissues, with increased amounts in certain disease states (Gaston, 1999).

Biochemical and cellular studies support the potential of GSNO to promote protein S-

glutathionylation. For example, GSNO reacts rapidly with the isolated proteins (e.g.,

papain, GAPDH, aldose reductase, creatine kinase, neurogranin, H-ras, c-Jun (Saavedra et al., 1997; Li et al., 2001; Mohr et al., 1999; Chandra et al., 1997; Giustarini et al.,

2005), and when rates were determined, this was shown to occur within minutes

(Giustarini et al., 2005; Mohr et al., 1999). In cells and cellular extracts, GSNO treatment has been shown to cause glutathionylation of carbonic anhydryase, H-ras, and

GAPDH (Ji et al., 1999; Mohr et al., 1999); and generation of endogenous NO via acetylcholine or bradykinin treatment was correlated with SERCA-SSG formation in arterial rings (Shelton et al., 2005). Taken together, these studies suggest that GSNO- mediated protein glutathionylation may be a feasible mechanism for protein-SSG formation in cells, although the relative rates of GSNO and protein-SSG formation in vivo need to be studied more directly in a redox signaling context to determine precursor-

29 product relationships. Furthermore, it is not well understood what protein properties or conditions favor protein-SNO vs. protein-SSG upon reaction of protein-SH with GSNO

(Giustarini et al., 2005; Park, 1988), and there is little information about the alternative reaction of protein-SNO with GSH to give protein-SSG (Scheme 1.1, p. 109, lower).

Thus, much remains to be examined concerning the convergence of ROS and RNS production with redox signaling and intermediate formation of protein-SSG.

Thiyl Radical Intermediate: Of the activated sulfhydryl intermediates, thiyl radicals

(RS) are among the shortest-lived (Wardman and von Sonntag, 1995); however, several lines of evidence support the likelihood of thiyl radical-mediated protein glutathionylation (and redox regulation) in vivo: (1) thiyl radicals are likely to be formed under redox signaling and oxidative stress conditions (Davies and Hawkins, 2000; Karoui et al., 1996; Kwak et al., 1995; Maples et al., 1990, Bradshaw et al., 1995; Davies et al.,

1997; Venters, Jr. et al., 1997; Cadenas, 1995); (2) decay of thiyl radicals via formation of disulfides (Scheme 1.1, p. 109, lower) is faster than other competing mechanisms of decay (Wardman and von Sonntag, 1995), (3) protein-SSG products are far more stable than thiyl radicals, making them more likely regulatory intermediates; and (4) diverse proteins undergo S-glutathionylation in the presence of GS-radical generating systems in vitro, with associated changes in protein function (Shelton et al., 2005). Also, there is evidence that this process may be enzyme-mediated (see Section 1.1.4A).

Thiosulfinate Intermediates: The thiosulfinate derivative of GSH (GS(O)SG) is a

- product of GSNO decomposition and is also formed by the the O2 /H2O2-producing system xanthine/xanthine oxidase (X/XO) (Li et al., 2001). Like sulfenic acids, thiosulfinates are highly reactive with thiols, forming disulfide and water (Huang and

Huang, 2002), suggesting both chemical and kinetic competence for glutathionylation of

proteins by reaction of GSH thiosulfinate with protein cysteines (Scheme 1.1, p. 109,

lower). This type of reaction was proposed for rat brain neurogranin (Li et al., 2001) and

for tyrosine hydroxylase (Sadidi et al., 2005), but in both cases it is uncertain whether

GS(O)SG or an alternative GSH derivative (e.g., GS or GSSG) was the primary mediator

of protein glutathionylation in situ. Moreover, in the case of tyrosine hydroxylase (Sadidi

et al., 2005), the authors presented evidence for sulfenic acid intermediates, so it is

possible that GSOH or protein-SOH played more important roles than GS(O)SG.

It is also feasible that protein thiosulfinates could react with GSH to form protein-

SSG and GSOH (Scheme 1.1, p. 109, lower). Indeed, Okamoto and coworkers. (2001) proposed that one mechanism of activation of matrix metalloproteinase precursors

(proMMPs) occurs via “S-glutathiolation … through glutathione disulfide S-oxide

GS(O)SR.” This suggestion is problematic, however, because biochemical characterization of the protein intermediates was inconsistent with thiosulfinate formation

(i.e., resistance to reduction by DTT (Hogg, 1979)), and no chemical or kinetic evidence was presented to support the supposition that MMP “thiosulfinate” proceeds to MMP-

SSG, Thus, much remains to be learned about the potential intermediacy of thiosulfinates in protein-SSG formation.

Sulfenylamide Intermediates: X-ray crystallographic analyses of PTP1B suggest that under extreme oxidative conditions, the sulfur and backbone nitrogen of active site

Cys215 form a cyclic, “sulfenylamide” bond (Salmeen et al., 2003) (van Montfort et al.,

2003). The PTP1B-sulfenylamide was fully reducible by GSH, suggesting PTP1B-SSG

as an intermediate (Salmeen et al., 2003). However, the kinetic competence of PTP1B-

sulfenylamide is uncertain. If sulfenylamide formation proceeds through a protein-SOH

intermediate as suggested (Salmeen et al., 2003; van Montfort et al., 2003), then it may not accumulate when GSH is present, because protein-SOH readily reacts with GSH to form protein-SSG (see above). Clearly, more must be learned about the rate of formation and stability of the PTP1B-sulfenylamide in the cellular milieu before its role as a signaling intermediate—and potential precursor to protein-SSG—can be assessed.

1.1.4 Catalysis of protein deglutathionylation

1.1.4A. Glutaredoxins

Glutaredoxin (Grx) is the most thoroughly characterized deglutathionylating enzyme, with the mammalian cytosolic form, i.e., Grx1, serving as the prototype (Mieyal et al.,

1995; Mieyal et al., 2008). Grx operates via a nucleophilic, double-displacement catalytic mechanism with both steps displaying high commitment to catalysis (Scheme

1.2A, p. 111, and Section 1.2). Rate enhancement by Grx1 has been attributed primarily to the unusually low pKa of the C22 thiol, which serves as the leaving group in the

second, rate-limiting step (Srinivasan et al., 1997).

Grx1 exhibits exclusive selectivity for protein-SSGs vs. other protein mixed disulfides (e.g., protein-SS-Cys and protein-SS-cysteamine, which are not substrates

(Gravina & Mieyal, 1993), and it is a far more efficient catalyst than other thiol-disulfide oxidoreductases for reducing protein-SSG (e.g., Grx1 displays a 5000-fold greater kcat/KM

for CSSG vs. Trx (Chrestensen et al., 2000)). Because it deglutathionylates diverse protein substrates (e.g., hemoglobin-SSG, papain-SSG, HIV-1-protease-SSG, actin-SSG

(Mieyal et al., 1995; Gravina and Mieyal, 1993; Davis et al., 1997; Wang et al., 2001b),

Grx is thought to operate as a general deglutathionylating enzyme; however, the

efficiency of Grx1-mediated deglutathionylation varies ~100-fold among protein-SSG substrates characterized thus far (Mieyal et al., 1995). The latter observation suggests that substrate preference may play a key role in cellular redox regulation by Grx1, and this could be documented by kinetic characterization of protein-SSG substrates that have already been implicated in physiological signaling (Adachi et al., 2004a; Adachi et al.,

2004b; Wang et al., 2003; Reynaert et al., 2006; Barrett et al., 1999a).

Grx1 appears to be responsible for most of the deglutathionylating activity in

mammalian cells, as shown by previous studies (Chrestensen et al., 2000; Jung and

Thomas, 1996); and this conclusion is corroborated by studies in which no

deglutathionylating activity (by standard assay (Chrestensen et al., 2000)) was detectable

in tissues from Grx1-knockout mice (Ho et al., 2007). The importance of Grx1-catalyzed

deglutathionylation in redox signaling in normal and pathological states is supported by

cellular studies implicating regulation by Grx1 in such diverse pathways as mitogenesis

(Barrett et al., 1999a), protein synthesis (Adachi et al., 2004a), cytoskeletal organization

(Wang et al., 2001b), Ca2+ homeostasis (Adachi et al., 2004b), and cell fate via deglutathionylation of PTP1B, Ras, actin, SERCA, and IκB kinase, respectively.

Recently, a second isoform of Grx (Grx2) was discovered, which exhibits key

similarities and differences compared with Grx1 (Gladyshev et al., 2001; Lundberg et al.,

2001) . Structurally, Grx2 exhibits <35% sequence homology with Grx1, but like Grx1,

it contains an active site CXXC motif (CSYC vs. CPYC for Grx1), Trx fold, and

glutathionyl stabilization site (Bushweller et al., 1994; Gladyshev et al., 2001; Yang et

al., 1998). Like Grx1, Grx2 exhibits deglutathionylating activity for peptide and protein

substrates, although its specific activity is 10-fold lower than that of Grx1 , (Lundberg et

al., 2001; Gladyshev et al., 2001). Chapter 3 describes studies that address the basis for

the lower specific activity of Grx2, and discusses the possible contribution of Grx2 to intracellular deglutathionylating activity.

The Grx isoform Grx5 is of unknown significance in mammals. It corresponds to the fifth Grx isoform discovered in S. cerevisiae, which has homologs in plant and human genomes. Whether Grx5 catalyzes protein deglutathionylation remains controversial.

First, there is disagreement whether yeast Grx5 displays activity with the pro-substrate hydroxyethyldisulfide (HEDS) (see also Section 1.2.6D) (Tamarit et al., 2003;

Rodriguez-Manzaneque et al., 1999; Shenton et al., 2002). Secondly, Shenton et al.

(2002) suggested that Grx5 functions as a deglutathionylase because its expression

correlates with deglutathionylation of a yeast GAPDH homolog Tdh3 in situ, but the

possibility of direct deglutathionylation of Tdh3 by Grx5 is questionable because its

subcellular localization is separate from Tdh3. Thirdly, it was reported (Tamarit et al.,

2003) that Grx5 deglutathionylates carbonic anhydrase in vitro, but the reaction was

performed at high [GSSG] without GSH, precluding the characteristic turnover

mechanism for Grx (see Scheme 1.2A, p. 111). Hence, the contribution of Grx5 as a deglutathionylating enzyme in mammals, or any organism, remains a frontier for future study (see also section 1.2.6D).

1.1.4B. Sulfiredoxin

Sulfiredoxin (SRx) is a small oxidoreductase, first characterized as catalyzing reduction of cysteine sulfinic acids formed within the active sites of 2Cys peroxiredoxins

(Biteau et al., 2003) (Table 1.1, p. 115). Recently, Findlay and colleagues (Findlay et al., 2006) proposed that SRx also acts as a deglutathionylating enzyme, based on

observations that overexpression of SRx in HEK293 cells diminished protein

glutathionylation induced by the diazeniumdiolate PABA/NO and that purified SRx

reversed PABA/NO/GSH-induced glutathionylation of actin and PTP1B. However, there

are several possible interpretations, including (1) SRx-mediated scavenging of the

reactive species that mediate protein-SSG formation, (2) reduction by SRx of an activated

intermediate form of proteins that are precursors of protein-SSG (e.g., protein-SOH), (3) blockade of sulfhydryl groups susceptible to glutathionylation by SRx via protein-protein interactions, and (4) deglutathionylation of protein-SSG by SRx.

While the first two possibilities were not tested directly, support for the third comes from the observation that SRx with a mutation in its active-site cysteine was still able to block PABA/NO-mediated protein glutathionylation, presumably via steric inhibition of glutathionylation sites. The likelihood of the fourth possibility, i.e., SRx-catalyzed deglutathionylation of protein-SSGs, is difficult to evaluate for several reasons. First, activity analyses of purified SRx towards glutathionylated proteins suggest that SRx acts as a highly selective deglutathionylating enzyme rather than one with broad specificity for protein-SSGs (like Grx, see Section 1.2.3, Step 1), and is thus unlikely to regulate global protein glutathionylation status. Specifically, Findlay and colleagues reported that purified SRx was shown to reverse PABA/NO/GSH-induced glutathionylation of actin and PTP1B (Findlay et al., 2006), with associated restoration of protein function in the case of PTP1B (previously shown to be inhibited by glutathionylation (Barrett et al.,

1999a; Barrett et al., 1999b)). Secondly, the effect of SRx-mediated diminution of

protein glutathionylation was demonstrated primarily in cells in which SRx was

overexpressed several-fold, calling into question the contribution of endogenously

expressed SRx to total intracellular deglutathionylating activity. Finally, tissues from

Grx1 knock-out mice provided by Dr. Ye-Shih Ho exhibit essentially no residual

deglutathionylating activity under standard assay conditions (Chrestensen et al., 2000), using bovine serum albumin-SSG (BSA-SSG) as the prototype substrate (Ho et al.,

2007), suggesting that other potential catalysts of protein-SSG deglutathionylation including SRx make little contribution to total intracellular protein deglutathionylating activity. Most recently selective deglutathionylation activity for SRx was documented.

Thus, Srx exhibited greater catalytic efficiency than Grx1 towards deglutathionylation of

2Cys Prx-SSG, but Srx did not catalyze reduction of prototype substrates BSA-SSG or

CSSG (Park, et al., 2009).

To further evaluate the likely contribution of SRx to physiological deglutathionylation in vivo, it will be critical to identify its physiological substrate(s) (in the presence of endogenous levels of enzyme) and to use the specific activity determined in in vitro experiments in combination with its intracellular concentration to assess its likely contribution to deglutathionylation activity in situ. Additional questions include its subcellular localization (in particular, whether it is in an oxidizing or reducing environment), and its contribution to deglutathionylation of protein-SSG intermediates following a physiological redox signaling stimulus (e.g., hormone or growth-factor- stimulated ROS production) vs. treatment with pharmacological agents such as

PABA/NO.

1.1.4C. Glutathione-S-Transferases

Several GSTs isoforms have been reported to catalyze protein deglutathionylation

(Garcera et al., 2006; Raghavachari et al., 1999). In each case, non-glutathionylated pro-

substrates (e.g., HEDS, S-sulfocysteine) were used to assess deglutathionylation activity.

To become glutathionyl mixed disulfides, these pro-substrates must first react with GSH,

forming βME-SSG or CSSG (Gravina and Mieyal, 1993; Mieyal et al., 1995).

Importantly, this assay cannot distinguish between catalysis of the glutathionylation step

(RSSG formation, characteristic of a GST-mediated conjugation) and the

deglutathionylation step (RSSG reduction, characteristic of a deglutathionylase such as

Grx). Indeed, Dal Monte et al. (1998) observed that bovine lens GSTμ catalyzes

formation of βME-SSG from HEDS and GSH but has a negligible effect on the rate of

βME-SSG deglutathionylation. Furthermore, when a protein-SSG substrate was used to

test GST-mediated deglutathionylation, addition of GST to a reaction mixture containing

Grx did not augment the deglutathionylation activity (Raghavachari et al., 1999) (for

further discussion, see (Gravina and Mieyal, 1993; Mieyal et al., 1995)).

1.1.5. Catalysis of Protein Glutathionylation

1.1.5A. GSTπ

Peroxiredoxins utilize a conserved active-site cysteines to reduce hydroperoxides to

alcohols (Rhee et al., 2005). A key step in catalysis is the formation of enzyme sulfenic

acid intermediates that either are reduced via a GSH-dependent mechanism involving Srx

(Park et al., 2009) (i.e., 1CysPRx) or form intramolecular disulfide bonds that are

typically reduced by TRx and/or TRx reductase (2CysPRxs) (Rhee et al., 2005).

Manevich et al. (2004) provided strong evidence that turnover of the 1CysPRx sulfenic acid intermediate is facilitated by GSTπ, which forms an interprotein complex and transfers GSH, converting 1CysPRx-SOH to 1CysPRx-SSG. The 1CysPRx glutathionyl mixed disulfide is then turned over by GSH (Scheme 1.3, p. 113). More recently, an

alternative mechanism was proposed for GSTπ-mediated reactivation of 1CysPRx in

which the 1CysPRx-SSG intermediate forms an interprotein disulfide with GST-SH

before being reduced nonenzymatically by GSH (Ralat et al., 2006). Although formation

of the intermolecular disulfide bond is chemically competent, its kinetic competence in

the catalytic cycle of peroxide reduction is questionable, involving multiple entropically

disfavored intermolecular disulfide exchange reactions.

The mechanism of delivery of GSH by GSTπ to effect conversion of a protein-SOH

to protein-SSG appears to be peculiar to its specific interaction with 1CysPRx.

Alternatively, Tew has suggested that GSTP (aka GSTπ) might promote more widespread

protein-SSG formation under certain conditions (Tew, 2007). This hypothesis seems to

be based on studies focused on the effects of PABA/NO treatment on various cell lines,

including the observation (Findlay et al., 2004) that PABA/NO-associated cytotoxicity

was somewhat diminished by knock-out of GSTπ and the report (Townsend et al., 2006)

that PABA/NO causes cytotoxicity, promotes protein-SSG accumulation, with kinase

activation and protein glutathionylation following similar time courses. On the basis of

these observations alone, it is difficult to link glutathionylation to cytotoxicity or to GSTπ

action. Therefore, further studies are necessary to determine whether GSTπ or other GST

isozymes promote glutathionylation of other proteins besides 1CysPRx.

1.1.5B. Grx

Oxidized derivatives of GSH increase during oxidative stress, including GS, GSNO,

and GSSG (Klatt and Lamas, 2000; Thomas et al., 1995), and these species are thought to contribute to formation of protein-SSG (see Section 1.1.2, above). Based on the low pKa

of C22 at the active site of Grx1 (Gravina and Mieyal, 1993) and the increased stability

of disulfide-anion radicals compared to thiyl radicals (Schoneich, 1995; Shoneich, 1995;

Wardman, 1995), we hypothesized that Grx1 might catalyze protein glutathionylation via

stabilization of the GS thiyl radical as an enzyme disulfide anion radical intermediate

(Grx1-SSG-), facilitating GS-radical recombination with a protein thiyl radical (Scheme

1.2D, p. 112). Indeed, both Grx1 and Grx2 promoted glutathionylation of reduced

protein substrates in the presence of a GS generating system (Gallogly et al., 2008;

Starke et al., 2003). It is plausible that this mechanism of glutathionylation is responsible

for p65-SSG formation and coincident inhibition of NFκB in a recent study of

hypoxia/N-acetylcysteine-induced apoptosis of pancreatic cancer cells (Qanungo et al.,

2007).

1.1.5C. Flavoprotein Sulfhydryl Oxidase (QSOX)

Sulfhydryl oxidase enzymes, ubiquitous in multicellular organisms but absent in

prokaryotes and yeast, utilize metals (metalloenzyme family) or flavins (flavoenzyme

family) to catalyze disulfide bond formation from diverse thiol substrates with

concomitant reduction of O2 to H2O2 (Thorpe et al., 2002) (Table 1.1, p. 115). Avian

and rat QSOX enzmes catalyze catalyze oxidation of low-molecular weight and protein

thiols. While the avian enzyme exhibits selectivity for protein sulfhydryls, forming

intramolecular disulfides, the rat enzyme oxidizes low molecular weight thiols (e.g.,

GSH) to intermolecular disulfides with higher efficiency (Coppock and Thorpe, 2006;

Ostrowski and Kistler, 1980).

Recognizing the ability of QSOX to utilize both low molecular weight and protein

thiols as substrates, the Mieyal research group has proposed that QSOX could catalyze

mixed disulfide formation between protein-SH and GSH, i.e., protein-SSG. To test this

hypothesis, David Starke (Mieyal laboratory) examined the ability of avian QSOX to

inactivate PTP1B, an established target of protein-glutathionylation in situ whose activity

is dependent upon maintenance of a reduced active site cysteine (Barrett et al., 1999a;

Barrett et al., 1999b). He observed QSOX- and GSH-dependent inhibition of PTP1B in a

time-dependent manner (Figure 1.1, p. 127; Gallogly and Mieyal, 2007), consistent with

glutathionylation of the active site Cys-215 of PTP1B (Barrett et al., 1999a).

Establishing QSOX as a glutathionylating enzyme will require further

characterization, including direct documentation of protein-SSG formation and

determination of catalytic efficiency. Evaluating the potential physiological role of

QSOX-mediated protein glutathionylation will require attention to subcellular localization, concentration, probable protein targets, and efficiency of glutathionylation under the conditions in which it is found intracellularly (i.e., the oxidizing compartments of the ER, Golgi, extracellular space (Thorpe et al., 2002)).

1.1.6 Other potential mechanisms of catalysis of protein S-glutathionylation

Other potential mechanisms of protein glutathionylation are based on analogy to the activities of enzymes known to catalyze similar reactions. For example, glutathionylation could occur through a glutathione peroxidase-like mechanism if protein-SH is substituted for one of the two GSH molecules in the typical reaction (Table 1.1, p. 115). Similarly, glutathionylation could proceed via a monooxygenase-like mechanism (utilizing either a heme-enzyme or a flavo-enzyme) as illustrated in Table 1.1, p. 115. To the best of our knowledge, potential glutathionylation activity of well-known peroxidases and monooxygenases has not been reported. Therefore it is not certain whether glutathionylation may be an important additional function of these enzymes, whether it is

instead catalyzed by undiscovered peroxidases or monooxygenases, or whether such

mechanisms of glutathionylation simply do not occur in cells.

1.1.7 Conclusions & Frontiers

Reversible protein glutathionylation plays critical roles in redox signal transduction,

as well as in the protection of cysteine sulfhydryls from irreversible oxidation. Grx1 is

the most extensively characterized deglutathionylating enzyme, and it appears to be

responsible for the majority of protein deglutathionylating activity in cells (Chrestensen

et al., 2000; Jung and Thomas, 1996; Steven A. Gravina, PhD thesis, Case Western

Reserve University, 1993; Ho et al., 2007). Recently, other candidate enzymes have been suggested to contribute to protein deglutathionylation intracellularly. Grx2, the mitochondrial Grx isoform, exhibits deglutathionylating activity in its monomeric form; however it is 10-fold less active than Grx1 (Lundberg et al., 2001; Gladyshev et al.,

2001), and its sequestration into inactive dimeric 2Fe2S clusters (Johansson et al., 2007;

Lillig et al., 2004) may interfere with efficient protein deglutathionylation in vivo (see also Chapters 3, 5.1). Grx5, a putative mitochondrial Grx isoform, also has been reported to exhibit deglutathionylating activity (Rodriguez-Manzaneque et al., 2002; Shenton et al., 2002; Tamarit et al., 2003), but it has not been evaluated under appropriate assay conditions or compared directly to the efficient Grx1. Preliminary studies on SRx suggested that it may function as a selective deglutathionylating enzyme (Findlay et al.,

2006), but it is unlikely to regulate global levels of protein glutathionylation. Indeed, the

proposed deglutathionylating activity of SRx towards diverse protein substrates should be

distinguished from alternatives; namely, prevention of glutathionylation, scavenging of

ROS/RNS, and reduction of precursors of protein-SSG. Recent further evaluation of SRx

has indicated efficient and selective catalysis of Prx-SSG deglutathionylation (Park et al.

2009). Analogous studies of other Grx isoforms, as well as potential discovery of other

deglutathionylating enzymes that may be highly specific for certain protein substrates,

will increase understanding of cellular mechanisms of protein deglutathionylation.

Identification of enzymes that catalyze protein glutathionylation is a major frontier in

the fields of redox regulation and signaling. Currently, there is evidence for selective

glutathionylation of a specific protein substrate by GSTπ via a catalytic mechanism

involving protein-protein interactions (Manevich et al., 2004; Ralat et al., 2006), but it is

unknown whether any GST isozyme catalyzes glutathionylation of other protein

substrates besides 1CysPRx, or whether other enzymes catalyze protein glutathionylation

with comparable selectivity. Preliminary evidence supports a potential role for

mammalian QSOX in catalyzing glutathionylation of proteins such as PTP1B (Gallogly

and Mieyal 2007, Figure 1.1, p. 127). However, characterization of the efficiency and

selectivity of QSOX-mediated protein glutathionylation, as well as documentation of

function in this manner by QSOX or related sulfhydryl oxidases in situ, represent areas

for future study. Finally, GSH peroxidase-like or monooxygenase-like enzymes, using

GSH and H2O2 or O2, respectively, as co-substrates, might catalyze glutathionylation of

proteins (Table 1.1, p. 115); however, there are currently no data to support this

hypothesis.

Future studies are likely to reveal new layers of complexity, including

characterization of more specific enzyme-substrate relationships and more diverse

mechanisms of catalysis and regulation, including compartmentalization of catalytic

proteins that mediate production and transfer of ROS, protein S-glutathionylation, and

protein-SSG reduction. As the frontier is advanced, continued attention to fundamental

principles of thermodynamic and kinetic competence under physiologically relevant

conditions need to be applied to interpretations of proposed reactions and reaction

intermediates representing intracellular events.

1.2 MECHANISTIC DETAILS OF CATALYSIS OF THIOL-DISULFIDE

EXCHANGE BY GLUTAREDOXIN

1.2.1 Introduction

As described in Section 1.1, Grx is the most specific and efficient catatlyst of

deglutathionylation in mammalian cells. This catalytic activity has been demonstrated to

regulate diverse cellular processes, including metabolism, calcium homeostasis,

cytoskeletal reorganization, hypertrophy, and inflammation (Mieyal et al., 2008). Kinetic

and mechanistic studies of Grx catalysis increase understanding of the bases for its

exquisite substrate selectivity and rate enhancement over uncatalyzed

deglutathionylation. They also provide information useful in future studies of the

enzyme, such as the strategic design of selective inhibitors.

1.2.2 Overall catalytic scheme for glutaredoxin

As described in Section 1.1, Grx-catalyzed deglutathionylation via a double-

displacement mechanism illustrated in Scheme 1.2A, p. 111. The mechanism was

elucidated from the results of 2-substrate kinetic analyses of hGrx1, in which 1/v vs. 1/[S]

plots yielded parallel lines for RSSG and GSH substrates (Gravina and Mieyal, 1993).

For such “ping-pong” catalytic mechanisms, KM and Vmax values for each substrate vary with the concentration of the second substrate, so “true” KM and Vmax values must be

determined by plotting 1/KM or 1/Vmax vs. 1/[S] and using the negative reciprocal of the each intercept (i.e., x-intercept for KM,true, y-intercept for Vmax,true) to determine intrinsic

kinetic constants. For hGrx1, these secondary plots produced a surprising result, namely

that the x and y intercepts projected to the origin (Gravina and Mieyal, 1993).

With respect to enzyme-substrate interactions, this result can be explained in one of

two ways. In the first, termed complete commitment to catalysis (Scheme 1.4A, p. 114),

k2 >> k-1, where k2 is the rate constant of the chemical step (the thiol exchange reaction) and k-1 is the dissociation of the enzyme-substrate complex. In this case, each enzyme- substrate binding event leads to a bond transformation reaction. The second explanation, termed an encounter reaction, describes a reaction between enzyme and substrate that occurs without formation of a reversible complex, so that no binding modes play a role in the reaction (Scheme 1.4B, p. 114). These possibilities can be distinguished via experiments using substrate analogs. In the case of complete commitment to catalysis, substrate analogs that bind tightly to the enzyme act as competitive inhibitors; for enzymes that operate via encounter reactions, substrate analogs are not inhibitory because they do not bind to the enzyme. The observation that S-methyl glutathione, which may

serve as a substrate analog for GSH or RSSG, does not inhibit hGrx1, even at millimolar

concentrations (Srinivasan et al., 1997), supports the interpretation that human

glutaredoxins operate via an encounter mechanism.

Catalysis of deglutathionylation by E. coli Grx1 appears to operate via an analogous ping-pong mechanism to human glutaredoxins (Bushweller et al., 1992). However, a recent report of the kinetics of yeast glutaredoxin 7 (ScGrx7, a monothiol glutaredoxin) displayed ping-pong kinetics, but non-zero intercepts were shown on secondary plots of

1/KM or 1/Vmax vs. 1/CSSG at three cysteinylglutathione (CSSG) concentrations

(Mesecke et al., 2008). It is possible that a more complete analysis involving more than

3 CSSG concentrations extending higher than the estimated KM might yield a result

corresponding to those for hGrx1 and hGrx2 (above); alternatively, this yeast Grx may be

dissimilar in this aspect of the double displacement mechanism, i.e., reversible ES

complexes may precede the covalent reactions.

1.2.3. Characteristics of individual steps of glutaredoxin catalysis of thiol-disulfide

exchange

Step 1: Reaction of the oxidized disulfide substrate with reduced Grx (Scheme

1.2A, Step 1, p. 111) – Step 1 represents a nucleophilic displacement reaction in which the catalytic cysteine thiolate within the Grx active site attacks the glutathionyl sulfur of a

protein-SSG mixed disulfide substrate, releasing protein-SH and forming a Grx-SSG

mixed disulfide intermediate. Glutaredoxin is highly selective for glutathionyl mixed

disulfide substrates, exhibiting no detectable reductase activity towards protein-SS-

cysteine substrates, which differ from protein-SSG by only 2 amino acids (Gravina and

Mieyal, 1993; Gallogly et al., 2008). Moreover, amounts of Grx far above those that are

effective for catalysis of deglutathionylation do not enhance the rate of glutathione

oxidation by hydrogen peroxide (glutathione peroxidase activity) (Starke et al., 2003;

Gallogly et al., 2008, Chapter 3).

Glutaredoxins exhibit selectivity not only for glutathionyl mixed disulfide substrates,

but also for the site of nucleophilic attack on those substrates. The oxidized product of

the first step of catalysis is exclusively the glutathionyl enzyme mixed disulfide (Grx-

SSG) as documented by mass spectrometric analysis on human and E. coli glutaredoxins

(Yang et al., 1998; Jao et al., 2006), and recent studies on E. coli Grx1 indicate that the orientation of the glutamyl moiety of the adducted GSH (i.e., the γ linkage) plays a crucial role in this specificity (Peltoniemi et al., 2006a; Saaranen et al., 2009).

The non-glutathionyl component of the disulfide substrate appears to be unrestricted.

Many protein-SSG and small molecule-SSG mixed disulfides (i.e., CSSG, and β- hydroxyethyl-SSG) are substrates (Mieyal et al., 1991b; Gravina and Mieyal, 1993;

Lundberg et al., 2001; Gladyshev et al., 2001; Johansson et al., 2004; Gallogly et al.,

2008). As discussed above, GSSG also serves as a substrate (Scheme 1.2B, p. 111), and this alternative substrate competition explains its apparent “inhibition” of E. coli Grx1 when added in increasing concentrations to a mixture containing tryptophanyl-peptide-

SSG (whose rate of deglutathionylation was measured fluorometrically) (Peltoniemi et al., 2006a). However, there are differences in reactivity among glutathionylated substrates when tested at the same fixed concentration of GSH (the reduced substrate).

These differences likely involve steric factors, as illustrated by comparison of met- vs. oxy-hemoglobin-SSG substrates (Gravina and Mieyal, 1993; Mieyal et al., 1995).

Following Step 1, wild-type Grx-SSG intermediate may undergo two disparate reactions: one leading to reduced Grx (Scheme 1.2A, step 2, p. 111), and the other to an intramolecular disulfide (oxidized) form of the enzyme (Scheme 1.2A, step 3, p. 111).

Step 2: Reaction of the Grx-SSG intermediate with a reduced thiol substrate

(Scheme 1.2B, Step 2, p. 111) – This is the rate determining step of the deglutathionylation reaction. For human Grx1, the pH rate profile of the Grx-catalyzed reaction matches that of deprotonation of the thiol on the reduced substrate, indicating the involvement of the reduced substrate thiolate in the rate-determining step (Srinivasan et

al., 1997; Gallogly et al., 2008). For Grx1, this overall maximal reaction rate for CSSG is ~2200 min-1 at 0.5mM GSH (Chrestensen et al., 2000); rates exhibited by mammalian

Grx2 isoforms are reported in Chapter 3.

Many thiols (including protein thiols) may be utilized as reduced substrates for Grx1-

SSG, and for most, the rate enhancement over non-catalyzed rates is ~1000-fold. This rate enhancement matches the predicted enhancement due to the difference in pKa of the

leaving group in the uncatalyzed reaction compared to the pKa of the Grx-thiolate as the

leaving group, according to the Bronsted theory (Srinivasan et al., 1997; Szajewski and

Whitesides, 1980). For GSH, there is an additional enhancement (~20-fold) beyond that

predicted by the leaving group effect, which suggests an enzyme-induced increase in nucleophilicity of the glutathionyl thiolate for the Grx-SSG intermediate (Srinivasan et al., 1997; Gallogly et al., 2008). To date, the basis for this enhanced nucleophilicity of

GSH is uncertain. Bronsted analysis of Grx1-SSG reduction by γ-Glu-Cys and Cys-Gly

(subsets of the GSH tripeptide γ-Glu-Cys-Cly) indicated enhancement of nucleophilicity of γ-Glu-Cys but not Cys-Gly (Srinivasan et al., 1997), implicating the γ-Glu moiety in the special interaction of GSH with Grx1-SSG. However, a recent comparison of Grx1- catalyzed reduction of peptide-SSG using GSH or Glu-Cys-Gly (ECG) as second substrates demonstrated no preference of hGrx1 for GSH over ECG (Saaranen et al.,

2009), suggesting that the γ-linkage of the Glu moiety does not confer special reactivity for Grx1-SSG with GSH. Clearly, further studies are required to verify the specific chemical moieties and interactions that underlie the special reactivity of GSH with hGrx1-SSG. By comparison, E. coli Grx1 exhibited faster reaction rates with GSH compared to ECG as a second substrate, leading the authors to conclude that the specific

orientation of the glutamyl moiety in GSH is more important for substrate selectivity in

E. coli than human Grx1.

As for RSSG, Grx1 exhibits an infinite KM,true towards GSH, indicating no reversible

substrate binding (as shown for RSSG in Step 1, (Gravina and Mieyal, 1993; Srinivasan

et al., 1997). This relationship is further supported by experiments in which incubation

of pig or human Grx1 with radiolabeled GSH yielded non-overlapping elution profiles of

protein and radioactivity in size-exclusion chromatography (Yang and Wells, 1991) . As

discussed above, millimolar concentrations of S-methylglutathione do not inhibit either

human Grx isoform, supporting an encounter-type mechanism for the second step of catalysis.

Side Reaction (Scheme 1.2B, p. 111, step 3): Formation of a Grx intramolecular disulfide from the Grx-SSG intermediate – An intramolecular reaction between the C-

terminal active site cysteine and the Grx-SSG intermediate can compete with turnover of the enzyme intermediate (Scheme 1.2B, Step 3, p. 111). The products are oxidized Grx

(i.e., Grx-S2) and reduced GSH. The intramolecular disulfide (Grx-S2) must react with 2

GSH to re-form the reduced enzyme for another cycle of catalysis, and this occurs

readily. That is, assays beginning with oxidized enzyme display full activity with no lag

phase, indicating that reduction of the intramolecular disulfide form of the enzyme occurs

at least as rapidly as the rate determining step (Mieyal et al., 1995).

Formation of Grx-S2 detracts from turnover of the Grx-SSG intermediate and is nonproductive. Therefore, to the extent it occurs, it is inhibitory. Removal of the C-terminal

active site cysteine by mutagenesis resulted in a 2-fold increase in specific activity for

Grx1 (Yang et al., 1998), indicating intramoleclar disulfide formation draws off about

50% of the Grx-SSG intermediate at steady state during catalysis. The analogous Grx2

mutant also exhibits an increased activity when compared to the unmutated enzyme

(Johansson et al., 2004; Gallogly et al., 2008; see Chapter 3). However, this phenomenon may not be generalizable to glutaredoxins from all species. For example, the E. coli Grx1 C14S mutant and analogous S. cerevisiae Grx2 C30S mutants are ~75%

less active than their wild-type counterparts towards small glutathionyl mixed disulfide

substrates (Bushweller et al., 1992; Peltoniemi et al., 2006a; Discola et al., 2008).

Importantly, it is not yet known whether this lesser activity relates directly to contribution

of the second cysteine to catalysis (i.e., a dithiol catalytic mechanism), or it is the result

of a mutation-induced conformational change that disfavors reactivity in some other way.

1.2.4 Evaluation of the mechanisms of the catalytic enhancement by glutaredoxin

In human Grx1, Cys 22 has been characterized as the active catalytic principle, and its

pKa (~3.5) (Mieyal et al., 1991b; Gan and Wells, 1987) has been shown to be responsible for the majority of the catalytic advantage of Grx over the nonenzymatic reaction

(Srinivasan et al., 1997). This can be explained by the observation that the second-order

rate constant of a thiol-disulfide exchange reaction increases by a factor of ~4 for each

one pH unit-decrease in the pKa of the leaving group (Gilbert, 1990). Thus, for Grx1, the

fold difference in rate constant of the catalyzed reaction (in which the leaving group in the rate-determining step is Grx1-SH, pKa ~3.5) vs. that of the uncatalyzed reaction (in

which the leaving group is BSA-SH, pKa ~8.5) is predicted to be 4ΔpKa, or 45 (~1000- fold) (Srinivasan et al., 1997). In contrast, thioredoxin (with an active site pKa of 6.7

(Kallis and Holmgren, 1980) exhibits very little deglutathionylating activity (Chrestensen et al., 2000).

The interaction of glutaredoxin’s catalytic cysteine with neighboring amino acids has been probed to understand the basis for thiolate stabilization reflecting the unusually low pKa. Examination of the NMR structure of the reduced enzyme suggested the nearby lysine (K19) might be responsible for the low pKa of Cys-22 (Jao et al., 2006; Sun et al.,

1998). According to this premise, computational studies in which K19 in human Grx1 was replaced by glutamine and leucine, and energy minimized, predicted that cysteine-22 of the mutated enzyme would have pKa values of 7.3 and 8.3, respectively. When actual

K19Q and K19L mutants were made in a form of the enzyme in which the three non- active site cysteines were mutated to serines (C7S, C78S, C82S), the resultant pKa values for the respective catalytic cyteines were both determined to be 3.7, i.e., little changed from wild type enzyme. This result indicated that the neighboring lysine cannot be solely responsible for the low C22 pKa and that a more complex set of interactions is ultimately responsible. Surprisingly, mutation of the C-terminal active site cysteine C25 resulted in about a one pH-unit increase in the C22 pKa, suggesting that the C25 thiol makes some contribution to the low pKa of the catalytic cysteine, either directly or conformationally.

Others have suggested multiple and varied interactions between the active cysteine and other amino acids on the protein, including H bonding within the active site (Foloppe and

Nilsson, 2007) and ion-dipole interactions with alpha-helix 2 (Jao et al., 2006; Kortemme and Creighton, 1995).

Importantly, the low pKa of glutaredoxin’s catalytic cysteine does not fully account for the observed rate enhancement in the presence of GSH. When GSH is used as the second substrate, second-order rate constants are further increased (by ~20-fold)

(Srinivasan et al., 1997) over rates using non-glutathionyl substrates. It appears that the

special rate enhancement of GSH can be attributed mainly to the γ-glutamylcysteine dipeptide subset of GSH, since γ-glutamylcysteine also confers an additional, albeit somewhat smaller, rate enhancement over non-GSH substrates, but cysteinylglycine (the other dipeptide subset of glutathione) does not. Although the basis for the differential enhancement of GSH nucleophilicity by human Grx isoforms is not yet known, the observation helps explain the difference in their specific activities. That is, the difference in catalytic cysteine pKa (~1 pH unit) accounts for only about half of the ten-fold lower

specific activity of Grx2 compared to Grx1. However, the additional rate enhancement in

the presence of GSH is 2.5-fold lower for Grx2 than Grx1 (8-fold vs. 20-fold), explaining

the remainder of the difference.

The basis for the distinct specific activities of dithiol glutaredoxins from S. cerevisiae

is not as well understood. Studies of purified, recombinant enzymes indicate a 15-fold higher specific activity for Grx2 vs. Grx1 (Discola et al., 2008; Luikenhuis et al., 1998).

Determination of the pKa of each enzyme’s catalytic cysteine revealed a difference of less than 1 pH unit, which was not sufficient to account for the 15-fold difference in specific activity (Discola et al., 2008). Potential differences in enhancement of glutathionyl nucleophilicity were not tested directly in this study; however, the authors used a combination of structural analysis and site-directed mutagenesis studies to propose a structural explanation for the higher activity of ScGrx2. Examination of the x-ray structure of ScGrx2 suggested that S23 and Q52 form a constellation of electrostatic interactions within the active site that appear to sequester the C-terminal active site cysteine (C30) away from the catalytic cysteine (C27). In contrast, C30 is oriented in close proximity to C27 in the published structure of ScGrx1 (Hakansson and Winther,

2007), possibly directed by alternative electrostatic interactions with residues A23 and

E52. Activity analyses of each enzyme’s C30S mutant support the structural data; that is,

ScGrx1 C30S exhibits higher activity than the wild-type enzyme (due to decreased

reaction between ScGrx1-SSG and the second cysteine, decreasing partitioning to the

side reaction), and ScGrx2 C30S has lower activity than wild-type enzyme. The authors

hypothesize that the sequestration of C27 in ScGrx2 (explained by interactions with S23

and Q52) prevents the enzyme from undergoing the nonproductive side reaction,

accounting for its higher activity vs. ScGrx1. In site directed mutagenesis experiments, swapping the residues implicated in C30 stabilization did increase the specific activity of

ScGrx1 and diminish that of ScGrx2, suggesting that they do account for part of the

specific activity difference between the two enzymes. It is important, however, to

address the authors’ hypothesis that the activity differences in the mutant enzymes can be

explained by altered partitioning to the side reaction. This could be done by comparing

the effect of the C30S mutation on enzymes in two backgrounds: wild-type and 23/52

mutant. Finally, since residue-swapping experiments only accounted for ~3-fold difference in activity between the two enzymes, other contributing mechanisms should be explored, including differences in enhancement of glutathionyl nucleophilicity.

1.2.5 Reports of complex formation between Grx and GSH

As described above for the catalytic Grx-SSG intermediate, there is evidence for interaction of the covalently bound glutathionyl moiety with specific residues of the Grx protein. NMR and x-ray crystallographic structures of several glutaredoxins in mixed disulfides with GSH (Bushweller et al., 1994; Yang et al., 1998; Nordstrand et al., 1999;

Yu et al., 2008; Discola et al., 2008; Hakansson and Winther, 2007) indicate that

different components of the GS-moiety are stabilized by specific residues on the Grx

molecule (Figure 1.4, p. 132). For human, yeast, and E. coli Grx-SSG structures, the

backbone of the Cys residue of GSH makes antiparallel, β-sheet-like H bond contacts with the backbone amide and carbonyl groups of a conserved Val. The carboxylate and animo groups of the γ-glutamyl moiety are stabilized by amino acids with complementary

charges in most Grx-SSG structures; however, these residues vary among the Grx

proteins of different species (see Figure 1.4, p. 132). The carboxylate of the glycyl moiety of the glutathionyl adduct is stabilized by H bond donors and/or a positively charged lysine in most Grx-SSGs; however, a recent study of hGrx1 in which the interacting Lys was mutated to Leu or Gln resulted in retention of glutathionyl substrate specificity (Jao et al., 2006), suggesting that this specific ionic interaction is not required

for reaction of Grx with the glutathionyl sulfur of the GS-containing mixed disulfide substrate. Moreover, analysis of the stability of mixed disulfides between E. coli Grx3 and various GSH analogs (Elgan and Berndt, 2008) suggested that the interaction between Grx and the glycyl group of the GS-moiety contributes little to stabilization of the glutathionyl moiety in the Grx-SSG mixed disulfide intermediate.

Although the kinetic data described above (i.e., encounter-type reaction mechanism

int with 1/KM approaching zero, and lack of potent inhibition by glutathionyl analogs,

Section I.A.2), indicate little or no affinity of Grx for reduced GSH, two recent reports

seem to challenge this interpretation. First, Lundberg et al. (2006) reported that some of the recombinant human Grx2 from a bacterial lysate could be purified via affinity chromatography with immobilized GSH-sepharose, although recombinant fusion proteins

(Grx2-β-galactosidase, or Grx2-GFP) could not. Secondly, analysis of an x-ray crystal

structure of reduced hGrx2 with reduced GSH led Johansson et al.(2007) to suggest that

Grx2 exhibits non-covalent, “high-affinity” binding to GSH. Specifically, co- crystallization of dimeric Grx2 under aerobic conditions in the presence of GSH resulted in monomeric Grx2 complexed with GSH. In this complex, the cysteine residues of the

Grx2 active site and GSH were not sufficiently close to represent a disulfide bond.

Accordingly, the cysteines were interpreted to be fully reduced, although a similar S-S

distance in the x-ray structure of yeast “Grx1p” (C30S)-SSG was interpreted to reflect

mono-oxidation of the mixed disulfide bond (Hakansson and Winther, 2007). The

majority of the Grx2-GSH interactions were similar to those of the Grx-SSG mixed

disulfides, with the glutathionyl Cys making antiparallel interactions with backbone

residues of a conserved Val, and the glycyl carboxylate forming an ion pair with a

conserved Lys (Figure 1.4, p. 132). However, the γ-Glu-Cys peptide bond was flipped

relative to previously published Grx-SSG structures, creating some unique H bonding interactions, and indicating a modified “binding” mode than that of a covalently bound

GS-moiety, although the same region of the Grx protein was occupied. Hence it is conceivable that the Grx-GSH complex detected in the crystal structure may represent a precursor to the (Grx2)2 2Fe2S complex.

The association of Grx2 with GSH was proposed to be “high-affinity” and inhibitory

towards deglutathionylation (Johansson et al., 2007), resulting in formation of a “dead- end complex” (Berndt et al., 2007). The main experimental evidence for this concept appears to come from site-directed mutagenesis studies in which mutation of T95 (which contributes several electrostatic interactions with the associated GSH) to Arg was associated with decreased Grx2 dimer stabilization by GSH, and higher

deglutathionylation activity towards βME-SSG. However, it was not confirmed whether the TR mutation resulted in decreased association of reduced Grx2 with GSH. In fact, other observations provide evidence against tight binding of GSH to Grx2. For example,

Grx2 is not inhibited by GSH analogs (Gallogly et al., 2008) Chapter 3), and removal of

GSH from solution results in rapid dissociation of the dimeric Grx2 2Fe2S complex

(Lillig et al., 2005; Berndt et al., 2007). The uncertainty regarding the strength of the potential Grx2●●●GSH noncovalent interaction would likely be resolved by measuring a dissociation constant directly. In fact, this approach was used by Noguera et al. to

estimate the relatively low affinity of P. tremula Grx C4 for GSH (Kd = 8.6 mM)

(Noguera et al., 2005).

1.2.6 Catalysis of Deglutathionylation by Other Glutaredoxins and Glutaredoxin

Domains

Glutaredoxins have been identified in most living organisms, from bacteria to

plants to mammals (Fernandes and Holmgren, 2004). Of these glutaredoxins, human

isoforms 1 and 2 are the best characterized kinetically (see above). Overviews of

glutaredoxin proteins from different species can be found elsewhere (Lillig et al., 2008;

Sagemark et al., 2007; Fernandes and Holmgren, 2004; Rouhier et al., 2004); here, we

review evidence for deglutathionylation activity in glutaredoxins from prototype

organisms (see Table 1.2, p. 117), with special emphasis on newly described Grx

proteins, and putative glutaredoxin domains on multidomain proteins.

1.2.6A. E. coli

E. coli contain 4 glutaredoxins, of which three (EcGrx 1-3) exhibit deglutathionylation activity towards protein, peptide, and/or small molecule glutathionyl

mixed disulfides (Peltoniemi et al., 2006a; Shi et al., 1999; Vlamis-Gardikas et al.,

1997). In a comparative study of glutaredoxins 1-3, E. coli Grx2 was shown to have the

highest activity towards βME-SSG (i.e., HEDS assay (Vlamis-Gardikas et al., 1997)),

accounting for an estimated 80% of total intracellular deglutathionylation activity

(Vlamis-Gardikas et al., 2002). Like human Grx1, E. coli glutaredoxins 1-3 exhibit

glutathionyl specificity in Step 1 of the deglutathionylation mechanism, with EcGrx1

forming exclusively EcGrx1-SSG following incubation with a glutathionylated peptide

(Peltoniemi et al., 2006a), and glutaredoxins 2 and 3 exhibiting no GSH-dependent

reductase activity towards insulin disulfide (Vlamis-Gardikas et al., 1997). Null mutant

E. coli strains for glutaredoxins 1, 2, or 3 are viable, although double mutant strains lacking EcGrx2 and EcGrx3 exhibit somewhat increased sensitivity to certain oxidants

(Vlamis-Gardikas et al., 2002). To our knowledge, a mechanistic link between decreased

deglutathionylation activity and increased oxidant sensitivity has not yet been

established.

EcGrx4 is the only monothiol glutaredoxin found in E. coli, and it exhibits

considerable similarity to yeast Grx5 (see below), including 37% sequence homology and

a monothiol, CGFS active site (Fernandes et al., 2005). Purified, recombinant EcGrx4

did not exhibit deglutathionylation activity in the HEDS assay (Fernandes et al., 2005),

nor did active site mutants of EcGrx4 containing a dithiol moiety (CGFC), or the

classical active site of human and EcGrx1 (CPYC), suggesting that a structural feature

outside of the monothiol active site may explain its lack of deglutathionylation activity.

When treated with GSSG, EcGrx4 forms an intramolecular disulfide (reduced by TR in

vitro), as well as a mixed disulfide with GSH (reduced by EcGrx1); however, whether

these disulfides form under native conditions—and whether EcGrx4 is a natural substrate

within the bacteria for EcGrx1 or—remains to be seen. Analysis of the NMR structure of

reduced EcGrx4 suggests that it may stabilize a covalently bound GS-moiety similarly to

EcGrx3 (Fladvad et al., 2005), but it is not yet known whether EcGrx4-SSG represents a catalytic intermediate, or simply a post-translational modification regulating other functions.

EcGrx4 appears to be required for cellular survival, since EcGrx4 knockout strains are not viable. There is some evidence that EcGrx4—like Grx5 of yeast and mammals— regulates iron homeostasis, since its expression is increased following Fe depletion

(Fernandes et al., 2005). It will be interesting to discover the role (if any) of the Grx active site in the critical functions of EcGrx4, given its incompatibility with catalysis of deglutathionylation.

1.2.6B. S. cerevisiae

To date, seven glutaredoxins have been identified in S. cerevisiae, and they have been divided into 3 groups according to structural and functional characteristics.

Glutaredoxins 1 and 2 (ScGrx1-2) are the only yeast glutaredoxins containing the classical CPYC active site motif. Analyses of lysates from mutant strains, as well as purified, recombinant enzymes, indicate that both ScGrx1 and ScGrx2 exhibit deglutathionylation activity, with ScGrx2 exhibiting approximately 15-fold higher specific activity compared to ScGrx1 (Luikenhuis et al., 1998; Discola et al., 2008).

Remarkably, mutation of the active site to CPYS has opposite effects on the activities of these two enzymes (Discola et al., 2008). Recently, the 20S proteasome was identified as a potential intracellular target for deglutathionylation by ScGrx2 (Silva et al., 2008).

Yeast glutaredoxins 3-5 were identified in a yeast genome sequencing project as a family of ORFs with homology to previously identified glutaredoxins, but containing non-classical, monothiol active sites (CGFS (Rodriguez-Manzaneque et al., 1999)).

Assays of lysates from null mutant strains suggest that all three glutaredoxins contribute to cellular deglutathionylation activity (i.e., decreased GSH-dependent reduction of βME-

SSG by the mutant strains), with the greatest contribution from ScGrx5. While ScGrx5 was also proposed to deglutathionylate Tdh3, the yeast homolog of GAPDH, in situ

(Shenton et al., 2002), studies of purified, recombinant enzyme exhibited little or no activity towards βME-SSG and a mixed disulfide between carbonic anhydrase III and

GSH (i.e., CAIII-SSG (Tamarit et al., 2003)). There are several potential explanations for this discrepancy, including inactivation of the enzyme during purification, removal of a cofactor from the cellular lysate, and an apparent lack of GSH in the assay of purified

ScGrx5 with CAIII-SSG (precluding turnover of the enzyme via the mechanism previously characterized for glutaredoxins, see above).

Although catalysis of GSH-disulfide oxidoreductase reactions by ScGrx5 remains uncertain, a catalytic mechanism of protein deglutathionylation was recently proposed

(Tamarit et al., 2003), involving both the active site and a non-active site cysteine

(C117). Support for such a mechanism came from mass spectrometric and HPLC analyses suggesting formation of a ScGrx5 intramolecular disulfide following incubation with GSSG, and also from the observation that a C117S mutant exhibits decreased deglutathionylation of CAIII-SSG. However, it appears that the deglutathionylation of

CAIII-SSG was assayed in the absence of GSH, and if GSH is involved in the rate determining step (i.e., reduction of the oxidized enzyme intermediate, as demonstrated

for human Grx1 and Grx2), then the relative deglutathionylation rates observed for WT

and C117S ScGrx5 may not represent a meaningful comparison. Thus, the

deglutathionylation capacity of ScGrx5 should be determined in a standard Grx assay

(i.e., in the presence of GSH), and compared to those of other Grx enzymes, to rigorously

test its deglutathionylation capacity.

ScGrx6-7 are the most recently described glutaredoxins in yeast, and are characterized by unusual active site motifs (CSYS and CPYS, respectively), as well as an apparent ability to form homodimers and/or -tetramers (Mesecke et al., 2008). Purified,

recombinant ScGrx6 and ScGrx7 exhibit deglutathionylation activity towards βME-SSG,

but do not reduce insulin intramolecular disulfide, indicating glutathionyl specificity in

the first step of the deglutathionylation reaction. Several lines of evidence indicate that

recombinant ScGrx6 is purified from E. coli as a GSH-stabilized, tetrameric iron-sulfur

cluster, which must dissociate to exhibit deglutathionylation activity (Mesecke et al.,

2008). As for hGrx2 (Lillig et al., 2005; Johansson et al., 2007), iron-sulfur cluster

formation appears incompatible with deglutathionylation activity for ScGrx6, raising the

important question of what percentage of endogenous ScGrx6 may exist in iron sulfur

clusters in vivo, and what stimuli trigger release of the active monomer.

Although the physiological roles of yeast glutaredoxins are not completely

understood, null mutant studies suggest that ScGrx1-5 serve antioxidant functions

(Luikenhuis et al., 1998; Rodriguez-Manzaneque et al., 1999), with some isoforms

exhibiting specificity for particular oxidative stimuli (e.g., ScGrx1, superoxide; ScGrx2,

H2O2 (Luikenhuis et al., 1998)). ScGrx3-5 appear to function in iron homeostasis, with

ScGrx3-4 implicated in the regulation of Aft1, a transcription factor regulating genes

59 involved in iron regulation (Ojeda et al., 2006), and ScGrx5 playing a critical role in iron sulfur cluster assembly (Muhlenhoff et al., 2003; Rodriguez-Manzaneque et al., 2002), also a likely function of its mammalian homolog (see below). For all of the yeast glutaredoxins, understanding the relationship between deglutathionylation activity and maintenance of redox balance and/or iron sulfur cluster homeostasis remains an exciting frontier for future study.

1.2.6C. Plants

Many genes encoding putative Grx enzymes have been identified in individual plant species (e.g., 31 in A. thaliana, (Rouhier et al., 2004)), but few biochemical characterizations of expressed Grx proteins have been reported. Grx from spinach

(Morell et al., 1995), rice (Sha et al., 1997) fern (Sundaram et al., 2008), and poplar

(Rouhier et al., 2001; Rouhier et al., 2002a; Rouhier et al., 2002b) exhibit activity towards the pro-substrate HEDS in assays containing GSH, GR, and NADPH. In contrast to human Grx enzymes, where mutation of the C-terminal cysteine in the active site increases activity, the analogous mutation of a poplar Grx decreases deglutathionylation activity by approximately two-thirds, suggesting that the side reaction involving Grx intramolecular disulfide formation (Scheme 1.2A, p. 111, Step 3) does not detract substantially from the catalytic rate for poplar Grx. Alternatively, this Grx may utilize primarily the dithiol mechanism for catalysis of deglutathionylation, or a deactivating conformational change accompanies the mutation.

In addition to deglutathionylation of βME-SSG, poplar Grx (along with GSH and GR) supports the peroxidase activity of a type C peroxiredoxin (Prx, (Rouhier et al., 2001)).

Although the proposed mechanism of coupling does not involve a Prx-SSG mixed

60 disulfide intermediate, further studies may indeed identify Prx-SSG as the substrate for

Grx in this coupled reaction

Poplar Grx C1 was the first glutaredoxin to be described as existing in a dimeric,

2Fe2S cluster (Feng et al., 2006), which is coordinated by one active-site cysteine sulfhydryl from each monomer, and two GSH molecules (Feng et al., 2006; Rouhier et al., 2007). As for hGrx2 (see above), the function of the cluster-coordinated poplar Grx dimer is not yet known, but functions in thiol-disulfide exchange and iron sulfur cluster assembly have been proposed (Rouhier et al., 2007).

1.2.6D. H. sapiens

hGrx5 –Several lines of evidence suggest that Grx5 isoforms from higher organisms share functional similarities with the enzyme from yeast (ScGrx5, above). For example,

(a) impaired oxidant defense in yeast mutants lacking ScGrx5 was rescued by expression of the chicken or human Grx5 genes (Molina-Navarro et al., 2006); (b) a zebrafish mutant lacking Grx5 exhibited defects in hemoglobin synthesis linked to decreased iron sulfur cluster biogenesis (Wingert et al., 2005), a function proposed for ScGrx5

(Muhlenhoff et al., 2003; Rodriguez-Manzaneque et al., 2002); and (c) molecular changes consistent with reduced FeS cluster synthesis were observed in red blood cells from a man with a genetic mutation lowering Grx5 mRNA to ~10% of control levels

(Camaschella et al., 2007). To our knowledge, neither isolated nor recombinant Grx5 from higher organisms has been assayed for deglutathionylation activity. Therefore, it is not yet known whether deglutathionylation activity contributes to any of the physiological roles identified to date for the enzyme.

TGR – Trx and GSSG Reductase (TGR or TR2) is a multidomain protein containing structural characteristics of TR (and GR), and Grx. Thus, the CXXXC motif, FAD- and

NADPH-binding domains, dimer interface domain and GCUG tetrapeptide are characteristic of TR protins. The N-terminal domain contains a CXXS motif (CPHS) and some residues conserved among glutaredoxins that have been implicated in stabilization of a covalently-bound glutathionyl moiety (see Section I.B) (Sun et al., 2001).

Both full-length TGR and the separate Grx domain are active in the HEDS assay (in which the pro-substrate HEDS is converted to βME-SSG following incubation with

GSH), indicating that the CPHS motif is a functional Grx active site. Whether the Grx domain of TGR utilizes the same catalytic mechanism for deglutathionylation as E. coli and human glutaredoxins has not been tested directly. It has been proposed that the selenocysteine (Sec, U) residue of TGR’s C-terminal CGUC motif was involved in the deglutathionylation mechanism (Sun et al., 2005). However, deglutathionylation activity of a TGR Sec  Cys mutant was measured in the absence of GSH, precluding formation of βME-SSG from HEDS. Therefore, reduction of βME-disulfide was measured rather than βME-SSG. It was also suggested that the Grx domain of TGR was responsible for the enzyme’s GSSG reductase activity, but this activity does not appear to have been tested with the Grx domain alone. Thus, whether the Grx domain of TGR is sufficient for the enzyme’s deglutathionylase and/or GR activities remains an open question and will require additional experimentation.

An additional catalytic activity recently reported for the Grx domain of TGR is protein disulfide isomerization, particularly involving mixed disulfide formation between glutathione peroxidase 4 (GPx4) and other proteins in developing sperm cells (Su et al.,

2005). Indeed, the Grx domain of TGR was active in protein disulfide isomerization

assays, and when immobilized on an affinity column, it formed DTT-reversible cross-

links with several proteins from sperm extract. Whether protein disulfide isomerization,

deglutathionylation, or Trx reductase activity represents the principal catalytic activity of

TGR in vivo—and how the magnitude of those activities compare to those of other thiol-

disulfide oxidoreductase enzymes—represent important questions in discerning the

physiological role(s) of this intriguing enzyme.

Variant 3 TR1 – A recent analysis of alternative splice forms of the thioredoxin

reductase 1 (TR1) gene revealed a specific isoform (named variant 3) containing an N-

terminal Grx domain similar to that of TGR, but with a CTRC active site motif (Su and

Gladyshev, 2004). Kinetic analyses of variant 3 TR1 revealed an unusual profile of

enzymatic activities. First, purified, recombinant variant 3 TR1 exhibited partial TR

activity; that is, it reduced DTNB, but it did not support reduction of insulin with Trx.

Removing the Grx domain restored insulin/Trx reductase activity, suggesting that the Grx

domain may interfere with the interaction between TR and TRx. Secondly, when tested

independently, the Grx domain of variant 3 TR1 was not active in the HEDS assay (i.e.,

GSH-dependent deglutathionylation of βME-SSG); however, an active site mutant

(CTRC  CPYC) exhibited some activity, suggesting that the unusual active site of the

Grx domain may preclude deglutathionylation activity. The Grx domain of variant 3 TR1 exhibited no dehydroascorbate reductase, GST, peroxidase, or protein disulfide isomerase activities, leaving the catalytic role of this domain unknown. The authors of this study suggest that full-length variant 3 TR1 may be a specific reductant for an unknown substrate, or possibly function as a protein disulfide isomerase.

PICOT – PKCθ-interacting cousin of thioredoxin (PICOT) was identified by a yeast-

2 hybrid screen for interacting partners of protein kinase θ (PKCθ) in T lymphocytes

(Witte et al., 2000). When overexpressed in human T cells, PICOT colocalized with

PKCθ, and inhibited JNK, AP-1, and NFκB activation. Structurally, PICOT consists of a

N-terminal “Trx” domain and 1-3 C-terminal repeats of a “PICOT” domain, depending upon the species of origin (Isakov et al., 2000). PICOT’s Trx domain exhibits 29% amino acid similarity to human Trx, but it is predicted not to exhibit Trx catalytic activity because it contains only one cysteine in its “active site” (APNC vs. CGPC in human Trx); thus, it has been proposed that PICOT may act as an antagonist of Trx in vivo (Witte et al., 2000).

Recently, the PICOT domain has been classified as a monothiol glutaredoxin domain (Lillig et al., 2008; Belli et al., 2002), apparently based on predicted 3- dimensional structure (Isakov et al., 2000) and sequence similarity to other gluratedoxins, including the CGFS active site motif, which is shared by some other recently described glutaredoxins (see above). To our knowledge, neither the catalytic activities (including deglutathionylation), nor the mechanisms of regulation of PKCθ signaling, have been demonstrated for PICOT. Understanding the capacity of PICOT to catalyze deglutathionylation would be particularly interesting in light of its association with PKC, as some PKC isoforms may be regulated by reversible glutathionylation (Shelton et al.,

2005).

1.3 REGULATION OF GLUTAREDOXIN ACTTVITY

1.3.1 Subcellular Localization and Effects on Grx Activity

Both the mechanisms and the consequences of dynamic regulation of Grx activity depend partially on its intracellular localization. Specifically, localization to subcellular compartments determines substrate availability, chemical environment, and proximity to signaling factors and intermediates that may affect enzymatic activity (e.g., kinases, reactive oxygen or nitrogen species; see below). Furthermore, distinct localization patterns among different tissues suggest different levels of activity (and capacity for regulation) within the organism. The mammalian Grx enzymes exhibit distinct subcellular localizations, related in some cases to localization sequences and/or differential splicing of the mRNA.

Grx1 is primarily a cytosolic enzyme, and it has been implicated in regulation via deglutathionylation of multiple cytosolic proteins, including PTP1B-SSG, Ras-SSG, actin-SSG, and procaspase 3-SSG (Barrett et al., 1999b; Adachi et al., 2004a; Wang et al., 2001b; Pan and Berk, 2007). Recently, Grx1 was shown also to reside in the intermembrane space of mitochondria isolated from rat tissues (Pai et al., 2007), although the mechanism of its mitochondrial localization is unknown. Immunohistochemical staining of endometrial tissue suggested nuclear localization of Grx1 (Stavreus-Evers et al., 2002), but this has not been confirmed by confocal microscopy, colocalization studies, or studies of isolated nuclei.

Three subforms of human Grx2 have been identified, each representing the product of alternative splicing of the gene’s five exons. Grx2a contains an N-terminal mitochondrial localization sequence, which is both necessary and sufficient to target Grx2 to the

65 mitochondria (Gladyshev et al., 2001; Lundberg et al., 2001; Enoksson et al., 2005).

Recently, Pai et al. (2007) documented endogenous Grx2 only in the matrix fraction of mitochondria isolated from rat heart and liver tissues, suggesting a specific intramitochondrial localization of Grx2a, separate from Grx1. mRNAs encoding Grx2b and Grx2c result from alternative splicing of a distinct first exon that does not encode a mitochondrial localization sequence (Lonn et al., 2008). When overexpressed in HeLa cells, both unconjugated Grx2 and GFP-Grx2 fusion proteins exhibit a diffuse staining pattern suggesting cytosolic and nuclear distribution. Unlike Grx2a, which appears to be expressed ubiquitously, mRNA encoding Grx2b and Grx2c was more restricted, detected only in cDNA libraries derived from normal testicular tissue or from certain immortalized cell lines. Levels of endogenous Grx2b and Grx2c within testes and in cancer cells have not been determined, but immunoperoxidase staining of testicular tissue sections with a nonspecific Grx2 antibody indicated cytosolic staining in spermatids, spermatogonia, and

Sertoli cells, suggesting the presence of endogenous Grx2b and/or Grx2c in normal testes. Both Grx2b and Grx2c exhibit deglutathionylation activities using the pro- substrate HEDS, and Grx2c forms a dimeric 2Fe2S cluster in vitro, although the physiological significance of this observation is not yet known.

Analogous findings were reported for five variants of mouse Grx2 (mGrx2) mRNA, encoding three subforms of the enzyme (Hudemann et al., 2009). mRNA encoding mGrx2a was present in most tissues, while testicular tissue contained the highest levels of alternative splice variants Grx2c and Grx2d. However, in contrast to human Grx2 variants, Grx2c and/or Grx2d expression may not be restricted to the testes among non- cancerous tissues. RT-PCR and colocalization studies suggest that these Grx2 subforms

may also be expressed in specific cell types within the stomach, spleen, and other tissues.

Isolated, recombinant mGrx2c (analogous to human Grx2c) exhibits activity towards

HEDS and can form a dimeric 2Fe2S cluster in vitro. Isolated, renatured Grx2d did not

exhibit these activities, but it is not clear whether this finding could be due to inactivation

during its resolubilzation from E. coli occlusion bodies, or to a potentially inhibitory domain coded by its additional exon (called IIIb).

Thus, the majority of non-cancerous mammalian cells (with the exception of testicular cells) are expected to contain Grx1 in the cytosol and mitochondrial intermembrane space; and Grx2 in the mitochondrial matrix. Cancerous and testicular cells may also express subforms of Grx2 in the cytosol and nucleus. These specific localizations of the

Grx isoforms not only determine the capacity for protein deglutathionylation within

specific subcellular compartments, but also the potential mechanisms of regulation of Grx

activity.

1.3.2. Enzyme concentration

1.3.2A Global concentration – In theory, the most straightforward way to modulate

Grx activity in situ is to change the content of active enzyme by altering rates of its

production or degradation. While little is known about the transcriptional regulation of

Grx1, some hormonal and metabolic stimuli are associated with increased protein content

and activity, including adriamycin treatment of MCF7 cells (Wells et al., 1995), estradiol

treatment of cardiomyoblasts (Urata et al., 2006), MPTP exposure in male mice

(Kenchappa et al., 2004), and culture of retinal Müller cells in high glucose media

(Shelton et al., 2007). Recently, Grx2 mRNA was found to be elevated in peripheral

blood mononuclear cells treated with the antitumor agent imexon (Baker et al., 2007),

presumably via a putative antioxidant response element (ARE) in its promoter sequence.

1.3.2B Local concentration – In the absence of stimuli, cytosolic Grx1 concentration

was determined to be ~1 µM in cytosol from human red blood cells (Mieyal et al.,

1991a), and similar concentrations are estimated from reports of Grx content in bovine

liver (Hatakeyama et al., 1984) and calf thymus (Luthman and Holmgren, 1982). In rat

liver mitochondria, Grx1 is estimated to be 0.1 μM in the intermembrane space, while

Grx2 concentration is about 1 μM within the matrix (Gallogly et al., 2008). In both

cases, calculations were performed on homogenized samples and represent an average

concentration of the entire subcellular compartment. However, it is conceivable the Grx

enzymes are maintained at higher local concentrations via association with structural

proteins (i.e., scaffolding) within specific intracellular compartments (e.g., plasma

membrane lipid rafts). Although we are not aware of direct evidence for this concept, it

may apply to regulation of actin glutathionylation in fibroblasts (discussed below).

1.3.3. Chemical environment/milieu

1.3.3A pH – Kinetic analyses provide insight into the influence of environmental factors on Grx activities in vivo. Deglutathionylation activity hGrx1 is pH-dependent, with an inflection point ~8.5 (Srinivasan et al., 1997; Gallogly et al., 2008) (see Section

1.2.3, Step 2). This pH dependence predicts variations in Grx activity among subcellular compartments, where pH varies considerably. For example, analyses utilizing pH- sensitive dyes suggest that the pH of the mitochondrial intermembrane space is close to

7.0 (Porcelli et al., 2005), in contrast to that of the cytosol (~ 7.4). Thus, calculations based on the pH-dependence of Grx1 activity predict that a single molecule of Grx1

would be expected to be about twice as active in the cytosol compared to the

intermembrane space. Of course, additional factors, such as differences in enzyme

concentration across subcellular compartments (see above), and localization of specific

substrates would also influence deglutathionylation activity in each of these

compartments. The pH-dependence of Grx2 activity will be addressed in Chapter 3.

1.3.3B GSH concentration – Kinetic characterization of hGrx1 predicts that differences in local GSH concentration will affect its deglutathionylation activity in situ.

That is, the parallel line patterns exhibited on double-reciprocal plots (Gravina and

Mieyal, 1993; Srinivasan et al., 1997; Gallogly et al., 2008) indicate that KM and Vmax

values for protein-SSG (i.e., the first substrate, Scheme 1.2A, p. 111) increase with

increasing concentrations of the second substrate (GSH). Many pathophysiological

stimuli alter GSH content and GSH:GSSG ratio (reviewed by Dalle-Donne et al., 2007);

so the dependence of Grx activity on GSH concentration appears quite relevant to the in

vivo condition.

1.3.3C. Reactive oxygen species (ROS) sources – Proximity to ROS could also regulate Grx activity, either via oxidative modification (see below), or by creating

reaction conditions that favor catalysis of thiyl radical scavenging or protein

glutathionylation. Thus, mitochondrial electron transport chain components as well as

•- • plasma membrane receptor-linked NADPH oxidases generate ROS (O2 , H2O2, OH),

which can lead to GS• formation. Human Grx1 scavenges GS•, or uses it as a substrate

for GS-transfer reactions, and we have investigated this activity also for hGrx2 (Starke et

al., 2003; Gallogly et al., 2008). Hence local production of ROS may favor catalysis of

glutathionylation by Grx. Indeed, Grx1 was implicated as the mediator of

glutathionylation of NFkB-p65 in pancreatic cancer cells under hypoxic conditions that

promote radical production (Qanungo et al., 2007).

1.3.4 Temporal Regulation of Glutaredoxin Activity

Grx-catalyzed deglutathionylation has been implicated in catalytic control of critical

processes relating to intracellular homeostasis and stress response, including cytoskeletal

reorganization (Wang et al., 2001b; Wang et al., 2003), mitogenic (Barrett et al., 1999a)

and hypertrophic signaling (Adachi et al., 2004a), inflammation (Reynaert et al., 2006;

Shelton et al., 2007), and Ca2+ homeostasis (Adachi et al., 2004b). Although there is

evidence that Grx activity itself is regulated both acutely and chronically in association with specific toxins, hormones, disease states, and extracellular stresses (below), the specific mechanisms by which Grx activity is regulated by these stimuli are not fully understood. The following section explores existing evidence for regulation of Grx activity in situ, and identifies future studies to elucidate the mechanisms further.

1.3.4A. Post-translational modification

Potential for phosphorylation – Reversible phosphorylation of serine, threonine, and tyrosine residues is a well-documented mechanism of enzyme regulation. To our knowledge, there is no evidence to date for regulation of Grx by phosphorylation; however, sequence analysis (Blom et al., 1999; Blom et al., 2004) of all three human Grx

isoforms reveals several putative phosphorylation sites, and identification of these sites

on published structures of reduced Grx1 (Sun et al., 1998) and Grx2 (Johansson et al.,

2007) suggests that they are solvent-exposed (Table 1.3, p. 120). Phosphorylation sites with the highest probability scores (i.e., 0.8-0.9) were found in Grx2, and included T41

(equivalent to T81 in the FASTA sequence (Lipman and Pearson, 1985)), immediately

following the CPYC active site motif, T97 (T137 FASTA), located on helix 4 adjacent to

the active site; and S118 (S158 FASTA). All three were consensus sites for protein

kinase C (PKC), and T97 also represents a generic kinase target. A high-probability, non-kinase specific phosphorylation site was also identified near the C terminus of Grx5

(S156 in its FASTA sequence).

None of the predicted phosphorylation sites appear to be conserved among the human

Grx isoforms (see Figure 1.3, p. 130), suggesting that phosphorylation could regulate the enzymes in an isozyme-specific manner. If phosphorylation is to be documented as a mode of regulation of Grx activity, then it will need to be demonstrated in situ, under physiological conditions, and associated with a functional change, analogous to the criteria for regulation by glutathionylation proposed by Shelton et al. (2005).

Oxidative modification – While Grx has been characterized as a regulator of protein thiol-disulfide status, some investigators suggest that its own cysteines may be modified by oxidation, with concomitant changes in activity (see below). Human Grx1 has 3 cysteines outside its active site (C7, C78, and C82). NMR structural analysis predicts they are all solvent-accessible (Sun et al., 1998), and thus susceptible to oxidative modification. In contrast, the two non-active site cysteines of Grx2 (C28, C100) appear to be involved in a structural disulfide bond (Johansson et al., 2007; Sagemark et al.,

2007). While these observations support the possibility of post-translational redox regulation of Grx, little is known about the specific modifications or their relevance to redox homeostasis in vivo.

Grx1 treated with HEDS is converted to multiple intramolecular and mixed disulfide forms according to mass spectrometric analysis, and fully active enzyme is regained by

71 adding it to the assay mix containing GSH (Papov et al., 1994). Likewise, both Grx1 and

Grx2 treated with GSSG exhibited a near total loss of thiol content, attributable to intramolecular and/or mixed disulfides by mass spectrometry (Hashemy et al., 2007), and this was associated with a 25% loss of activity for Grx1 that could be fully reversed by

DTT. The in vivo relevance of this relatively small deactivation by GSSG is questionable. First, the oxidative challenge (10 mM GSSG in the absence of GSH) does not represent a physiological condition. Secondly, the absence of GSH may allow oxidations that would not occur in its presence, resulting in “false positive” modifications. These considerations are useful in interpreting the results of additional oxidation experiments (below).

i. H2O2 – Human Grx1 and Grx2 undergo partial inactivation at millimolar concentrations of H2O2 in vitro, with Grx1 being slightly more sensitive to inactivation

(Starke et al., 1997; Gladyshev et al., 2001; Qiao et al., 2001; Hashemy et al., 2007).

H2O2-treated Grx1 could be reactivated by DTT, GSH, or the Trx/TR system (Qiao et al.,

2001). In contrast, high concentrations of GSH (5mM) but not low GSH concentrations

(0.5mM), DTT, or the Trx/TR system restored Grx2 activity following H2O2 treatment

(Gladyshev et al., 2001). The potential effect of H2O2 on Grx activity is an important consideration since H2O2 may reach relatively high concentrations in subcellular compartments where Grx isoforms are found. It will be critical to document the dose- dependence of H2O2-mediated inactivation in the presence of GSH since physiological levels of GSH may offset the oxidative effects of H2O2 on the enzymes, thus diminishing the relevance of the modifications.

• ii. OH – Hydroxyl radicals are formed by reaction of H2O2 with metal ions such as

Fe2+ and Cu+ (Halliwell, 1996), and are widely implicated in oxidative damage in disease

states such as ischemia-reperfusion injury. Starke et al. (1997) observed inactivation of

Grx1 in a dose-dependent manner by a hydroxyl radical generating system in vitro.

However, stop-flow ischemia-reperfusion of intact rabbit heart did not affect Grx1 activity, suggesting either that ischemia-reperfusion did not cause sufficient oxidative stress to inhibit the enzyme, or that the inhibition was reversed by a component absent from in vitro assays (e.g., GSH, see above).

iii. Reactive nitrogen species (RNS) – Reactive nitrogen species, including

- •- peroxynitrite (ONOO , a product of O2 and NO), and S-nitrosoglutathione (GSNO) are produced under redox signaling and oxidative stress conditions (reviewed in (Gallogly and Mieyal, 2007)) and are associated with both glutathionylation (Adachi et al., 2004b;

Clavreul et al., 2006a) and nitrosylation (Klatt and Lamas, 2000; Mohr et al., 1999;

Giustarini et al., 2005) of protein cysteines. To date, it appears that mammalian Grx

isozymes exhibit distinct sensitivities to RNS. For example, rat liver Grx1 (Aykac-Toker

et al., 2001) and recombinant Grx1 (Hashemy et al., 2007) were largely inactivated following ONOO- exposure in vitro, while Grx2 activity was not affected (Hashemy et al., 2007). The ONOO--dependent inactivation of the recombinant Grx1 was not

reversed by DTT, suggesting a non-disulfide modification (cysteine sulfinic or sulfonic

acid formation, or tyrosine nitration). The lack of reversal precludes this modification as

a regulatory mechanism. For GSNO, the inhibition of Grx1 was influenced by O2, with

greater inhibition and refractoriness to reversal under anaerobic conditions (Hashemy et

al., 2007). The GSNO-mediated inhibition correlated with appearance of an absorption

band characteristic of protein-SNO, but a specific site of Grx1 nitrosylation was not

determined. Monomeric Grx2, like Grx1, exhibited a near total loss of thiol content upon

exposure to GSNO; however, Grx2 activity was not affected by treatment. Dimeric Grx2

dissociated into active monomers in the presence of GSNO, suggesting a potential

mechanism for regulation in situ (Hashemy et al., 2007).

Proteolysis – A recent study in lung epithelial cells suggests that Grx1 can be

degraded by caspases following stimulation of the cells with Fas ligand (Anathy et al.,

2009). Specifically, Fas ligand treatment led to a diminution of Grx1 content and

activity, which was prevented by pretreating the cells with a caspase inhibitor.

Incubation of purified Grx1 with activated caspase 3 or 8 produced an 8 kDa fragment that was immunoreactive with an anti-Grx1 antibody, suggesting that the diminution of

Grx1 observed in Fas-treated cells may result from caspase-mediated cleavage of the

enzyme. To our knowledge, this is the first demonstration that Grx1 activity may be

regulated by protein degradation. Moreover, this work represents an intriguing contrast to that of Wang et al. (2007), who identified caspase 3-SSG as a substrate for Grx1

(Wang et al., 2007; see Section 1.3.4C). Certainly, further studies will increase

understanding of the enzyme-substrate relationship between Grx1 and caspase 3.

Many splice variants of human Grx2 have been identified (Lonn et al., 2008;

Lundberg et al., 2001); and one (termed Grx2a) codes for an N-terminal mitochondrial

signal sequence. The functionality of the signal sequence was confirmed by

demonstrating that Grx2-GFP fusion proteins were targeted to the mitochondria

(Lundberg et al., 2001; Gladyshev et al., 2001), and that immunoreactive Grx2 was

isolated from the matrix fraction of mitochondria from rat tissues (Pai et al., 2007). The

mature, truncated form of Grx2 is about 4 times more active than the full-length form,

possibly due to steric interference by the signal sequence (Lundberg et al., 2001;

Johansson et al., 2007).

Analysis of the predicted sequence of human Grx5 using using ProP1.0 software

(Duckert et al., 2004) reveals two putative cleavage sites for proprotein convertase (PC).

PC enzymes cleave pre-pro-proteins at specific sites containing single or paired basic

amino acids, often prior to secretion. No PC cleavage site was detected for Grx1,

although it is speculated to be a secreted protein (Lundberg et al., 2004; Peltoniemi et al.,

2006b).

Metal coordination – Cd2+ is a non-redox active, toxic heavy metal ion that

characteristically binds to vicinal dithiols, and Cd2+ inhibits Grx1 in a dose-dependent

manner in vitro, apparently by coordinating to the thiolate forms of the neighboring

active site cysteine moieties (Chrestensen et al., 2000). In HT4 cells, Cd2+ treatment was

correlated with increased protein-SSG content (Figueiredo-Pereira et al., 1998); and

Cd2+dose-dependent inhibition of deglutathionylation activity was observed in Jurkat and

H9 cells (Chrestensen et al., 2000). In contrast, a mutant strain of S. cerevisiae lacking

yeast Grx2 exhibited decreased protein-SSG content following Cd2+ exposure (Gomes et

al., 2008), suggesting that yeast Grx2 may exhibit primarily glutathionylating activity in

situ, as observed for the human Grx enzymes in vitro under glutathionyl radical

generating conditions (Gallogly et al., 2008; Starke et al., 2003) . Since Cd2+ is not likely

available to coordinate to the Grx enzymes naturally, and confers an essentially

irreversible inactivation (Chrestensen et al., 2000), this does not represent a likely

mechanism of regulation.

Recent studies on human Grx2 (Lillig et al., 2005; Berndt et al., 2007; Johansson et

al., 2007) and poplar GrxC1 (Feng et al., 2006; Rouhier et al., 2007) suggest that these

isozymes may exist naturally within dimeric 2Fe2S clusters, and that enzyme activity

may be directly influenced by dimer integrity. Structural analysis of recombinant Grx

from both species suggests that coordination to the 2Fe2S center utilizes 2 sulfhydryl

groups per monomer: one from the catalytic cysteine, and the other from a non-

covalently associated GSH molecule (Johansson et al., 2007; Feng et al., 2006; Rouhier

et al., 2007). Because the iron-coordinated cysteine thiolate is not available for nucleophilic attack (i.e.,it is complexed to the positively charged metal ion), it is not

surprising that the Grx2 dimer exhibits no detectable catalytic activity (Lillig et al.,

2005). In HeLa and BL30 cells cultured with 55Fe, the majority of immunoprecipitated

Grx2 was radioactive, indicating association with the radiolabeled iron and suggesting

that Grx2 may be predominantly dimerized (i.e., inactive) under nonstressed conditions in these cells. Clearly, identifying factors that lead to the release of active enzyme is critical for predicting Grx2 activity within the mitochondrial matrix. In vitro, a variety of thiol

oxidants (e.g., GSSG (Lillig et al., 2005)), reductants (e.g., dithionite, ascorbate (Lillig et

al., 2005)), and RNS (e.g., GSNO, SIN-1 (Hashemy et al., 2007)) disrupt the Grx2 dimer, but intracellular regulators of dimer status remain unknown. Finally, it has been suggested that Grx2 may function to regulate iron sulfur cluster homeostasis (Lillig et al.,

2005), but evidence supporting such a role—including whether its thioltransferase activity is required—represents a direction for future investigation.

1.3.4B Protein-protein interactions

Apoptosis signal-regulating kinase (ASK1) – Association of Grx1 with ASK1 has beeen proposed to regulate ASK1 activity—and subsequently apoptosis—in MCF7/ADR cells subjected to metabolic stress (Song et al., 2002). Specifically, the ability to co- precipitate endogenous Grx1 and ASK1 was lost upon glucose deprivation, and the dissociation of the Grx1-ASK complex was associated with ASK1 activation and increased apoptosis. The observation that Grx1-ASK1 dissociation was blocked by mutation of either or both of Grx1’s active site cysteines led the authors to propose that fully reduced Grx1 is the form that associates with ASK1, and formation of the active site intramolecular disulfide of Grx1 is required for its dissociation from the heterodimer

(Figure 1.4A, p. 132). If the heterodimerized Grx1 is truly in its reduced form, then it should be catalytically competent; however, it is possible that the site of interaction with

ASK1 (or the subcellular localization of ASK1) could restrict its availability to protein-

SSG substrates. On the other hand, the released Grx1-disulfide should be quickly reduced by GSH so long as GSH is not depleted. Clearly, testing cytosolic Grx1 activity before and after glucose deprivation would help answer this question. The result of such experiments would be particularly interesting given the apparently opposing effects of

Grx1 on apoptotic signaling (i.e., Grx1 overexpression protects H9c2 cells from apoptosis via regulation of Akt activity (Murata et al., 2003), but Grx1-catalyzed deglutathionylation of caspase-3 supports apoptosis in TNFα-treated endothelial cells, see below (Pan and Berk, 2007). Thus, the increase in ASK1 activity attributed to its dissociation from Grx1 could be either potentiated or countered by the increase in Grx1 activity following its release.

Also to be considered is the percentage of total Grx1 in complex with ASK1 under

resting conditions. The heterodimerization status of Grx1, if inactive in this bound form,

would determine the deglutathionylase activity available for cytosolic protein-SSG

targets. Resolving these important questions warrants further study.

Pro-caspase-3 – Another protein that appears to co-localize with Grx1 is pro-caspase-

3. Immunoprecipitation of Grx1 from bovine aortic endothelial cells pulled down a

protein that reacts with an anti-caspase-3 antibody. This interaction was almost

completely lost following TNF-α/cycloheximide treatment, leading Pan and Berk (2007)

to propose a model shown in Figure 1.4B, p. 132. That is, Grx1 and caspase-3 are

associated under resting conditions, but TNF-α stimulation causes Grx1 to

deglutathionylate pro-caspase 3-SSG, leading to its cleavage and activation, which

promotes downstream apoptotic events. As discussed below the specific mechanism of

Grx1 activation by TNF-α has not yet been identified.

1.3.4.C Examples of apparent rapid activation of Grx

FGF-induced actin-SSG Deglutathionylation (Figure 1.5A, p. 134) – Treatment of

NIH 3T3 cells with FGF resulted in robust deglutathionylation of β-actin within minutes

(Wang et al., 2003). While steady-state levels of actin-SSG were not sensitive to

manipulation of Grx1 content, its FGF-induced deglutathionylation was completely

blocked by knockout of Grx1 by stable transfection of siRNA targeting Grx1. In

addition, actin-SSG status was correlated inversely to actin polymerization (Wang et al.,

2001b) and cytoskeletal reorganization (Wang et al., 2003), suggesting that FGF-induced actin deglutathionylation may aid cytoskeletal changes involved in the proliferation response. While these observations could be explained by acute activation of Grx1,

several conceivable explanations need to be distinguished, including: (a) activation of

Grx1 deglutathionylation activity (via post-translational modification, conformational

change, or some other mechanism), (b) exposure of the actin-SSG disulfide bond to

already active Grx1 (via conformational change in actin or other cytoskeletal components), and (c) rearrangement of Grx1 and actin-SSG into close proximity via

scaffolding or protein-protein interactions. Experiments that would address these

possibilities include testing the effect of FGF treatment on global Grx1 activity (i.e.,

distinguishing changes in activity from changes in substrate availability), investigating

the subcellular localization of Grx1 before and after FGF stimulus, and performing pull-

down experiments with antibodies to actin and Grx to determine whether the two proteins

may be co-localized through mutual protein-protein interactions.

Shear stress (Figure 1.5B, p.134) – Flow, or fluid shear stress, represents the

frictional force of blood acting on the surface of vascular endothelial cells (Wang et al.,

2007). Disruption of physiological flow rates appears to alter signal transduction and

gene expression patterns in endothelial cells, although the specific mechanisms are not

yet fully elucidated (Chiu et al., 2008). Wang et al. (2007) observed that cultured bovine

aortic endothelial cells exhibited a brief (5-10 minutes) but robust increase in total Grx

activity upon exposure to shear stress. Grx activation was correlated to increases in Akt

and eNOS phosphorylation, and it was blocked by siRNA directed to Grx1. This short

time course precludes induction of the enzyme and suggests an acute, reversible

mechanism of regulation of Grx1. The authors propose that the increase in Grx1 activity

is explained by a similarly acute increase in the activity of GSSG Reductase (GR), the

enzyme that reduces GSSG, the second product of Grx-mediated protein

deglutathionylation (Figure 1.5B, p.134). This model is based on observations that shear

stress also resulted in increased GR activity, and BCNU (a GR inhibitor) blocked the

downstream effects of shear stress-induced Grx1 activity. Some assumptions related to

this interpretation should be tested explicitly, namely that the BCNU did not inhibit Grx1

directly; and that GR activity is limiting in this system (i.e., that an increase in Grx

activity can only be “realized” if GR activity is increased). If GR activity is not limiting,

then future studies should focus on events such as post-translational modification of

Grx1, etc. (see above).

TNF-α-induced caspase-3 deglutathionylation (Figure 1.5C, p. 134) – As discussed above, treatment of bovine aortic endothelial cells with TNF-α and cycloheximide (CHX) resulted in an increase in total Grx activity (Pan & Berk, 2007). Although the time course of activation by TNF-α/CHX (3-6 hours) is longer than those observed for FGF and shear stress exposures, it is unlikely to be explained by changes in protein expression or degradation because the time course is much shorter than the half life of the enzyme

(Grx1 t1/2 ~ 1.5 days (Wang et al., 2003)). Since Grx1 and its substrate (pro-caspase 3-

SSG) appear to be physically associated at baseline (see above), a likely mechanism of

activation would involve a post-translational modification or conformational change allowing active Grx1 to access the glutathionylated cysteine of the pro-caspase 3 substrate.

1.3.5 Conclusion

The functional consequences of the deglutathionating activity of the glutaredoxins are likely profound. The high catalytic rate and rate enhancement over the low uncatalyzed reaction rate imply that Grx controls the glutathionylation status of proteins

that are specifically glutathionylated, analogously to the action of phosphatases. The

specificity of Grx for protein-SSG substrates combined with the enhanced reactivity of

Grx-SSG with reduced GSH suggest that reversible glutathionylation is an organized and

systematic signaling system, analogous to the kinase/phosphatase signaling cascades.

Modulation of intracellular Grx activity by hormones, stress factors, and disease states

suggests an additional level of regulation of protein glutathionylation status via Grx in

vivo. All of the biochemical features that this catalytic system exhibits may uniquely

poise glutaredoxin as a sensitive and important cellular effector.

1.4 GLUTATHIONYLATION AND GLUTAREDOXINS IN

CARDIOVASCULAR DISEASE

1.4.1 Cardiovascular Diseases and Alterations in Protein-S-glutathionylation Status

Within the cardiovascular system, protein S-glutathionylation is emerging as a critical

signaling mechanism and consequence of oxidative insult, such as ischemia/reperfusion

injury (Eaton et al., 2002, see below). Protein S-glutathionylation regulates numerous physiological processes that are important in cardiovascular homeostasis and/or perturbed in disease, including myocyte contraction, oxidative phosphorylation, protein synthesis, vasodilation, glycolytic metabolism, and response to insulin (summarized in Table 1.4, p.

122). This section discusses evidence that perturbations in protein glutathionylation status, and/or regulation of Grx activity,contribute to the etiology of cardiovascular diseases such as myocardial infarction, cardiac hypertrophy, and atherosclerosis.

1.4.2 Myocardial Infarction

Eaton and colleagues (2002) analyzed the effects of cardiac ischemia-reperfusion on

glutathionylation of the cardiac proteome. Isolated rat hearts were perfused with

biotinylated GSH following stop-flow ischemia, and glutathionylated proteins were

detected via blotting with streptavidin-HRP. According to this analysis, overall protein

glutathionylation was increased approximately 15-fold following ischemia-reperfusion

(IR), with the majority of the glutathionylation events occurring early in the reperfusion

period. Some caution is required in interpreting these results because trapping the

modified proteins as protein-SSG-biotin may inhibit their deglutathionylation and

overestimate the degree of glutathionylation that would occur otherwise (Shelton et al.,

2005).

In the same study, GAPDH was identified as a prominent cardiac protein

glutathionylated during IR. GAPDH immunopurified from ischemic tissue exhibited

DTT-reversible loss of function, suggesting that GAPDH glutathionylation is likely

inhibitory in vivo. While the consequences of GAPDH inhibition on cardiac function were not explored in this study, logical possibilities include: (1) contribution to the blockade of glycolysis characteristic of ischemic injury, (2) interference with translocation to the nucleus, resulting in increased apoptosis (Ishitani et al., 2003; Colell

et al., 2007), or (3) little to no effect, with the modified cysteine of GAPDH, serving

primarily as a “sink” (or marker) for excess oxidants rather than a site of homeostatic

regulation. Importantly, GAPDH activity was restored by the end of the reperfusion

period, suggesting that for this protein at least, glutathionylation may serve as a

temporary modification to protect catalytic cysteines from irreversible oxidation.

Evidence for actin glutathionylation was demonstrated in a rat model of in vivo IR

(Chen and Ogut, 2006). Homogenates of ischemic tissue subjected to Western blot analysis with an anti-GSH antibody exhibited a prominent band corresponding to the molecular weight of actin, and immunoprecipitated actin reacted with the same antibody in a DTT-reversible manner. Studies on isolated G-actin indicated that glutathionylation delayed its rate of polymerization (consistent with a previous study on the effect of actin glutathionylation in A431 cells (Wang et al., 2001b)), and decreased the cooperativity of its binding to tropomyosin, suggesting that actin-SSG formation contributes to the decline in cardiac contractility observed during ischemia. This interpretation could be strengthened by determining the glutathionylation status of actin following reperfusion.

Since contractility is generally recovered in this model by the end of the reperfusion period, it would be expected that actin-SSG levels would decline with a similar time

course, providing the basis for the improved contractility following IR insult.

In contrast to actin and GAPDH, mitochondrial complex II exhibits the opposite

glutathionylation pattern following IR, i.e., de-glutathionylation. In vivo IR, as well as

stop-flow ischemia of isolated rat heart, resulted in decreased immunoreactivity of the 70

kDa subunit of complex II (i.e., semiquinone reductase, SQR) with an anti-GSH antibody

(Chen et al., 2007). Studies of isolated SQR indicated that glutathionylation increased

- electron transfer activity somewhat and decreased leakage of superoxide (O2 ), suggesting that IR-associated deglutathionylation contributes to the decrease in SQR

function observed during IR.

What could explain the divergence between the glutathionylation pattern of the

general cardiac proteome (including actin and GAPDH) and that of SQR during IR?

Possible contributing factors include the accessibility and intrinsic reactivities of

modified cysteines, their proximity to sites of ROS production (as well as to Grx), and

structural features that may stabilize the glutathionylated—or thiolate—status of the

modified cysteine. Overall, the divergent results of these studies reinforce the concept

that a single oxidative stimulus (e.g., IR) can affect glutathionylation status of different

protein cysteines in different directions. Moreover, understanding the basis for these

differences will require a greater understanding of the regulation of factors influencing

protein glutathionylation status (e.g., concentrations of ROS, Grx1, and Grx2) within

specific intracellular compartments, as well as quantitative relationships among protein-

SSG events, the magnitude of alteration in protein activity, and resulting impact on cellular function.

Understanding the role of protein glutathionylation in myocardial infarction also can be approached by manipulating Grx, the primary intracellular deglutathionylating enzyme

(Chrestensen et al., 2000; Jung and Thomas, 1996). To this end, mouse models with embryonic knockout of Grx1, as well as overexpression of Grx1 and Grx2 transgenes, were developed and subjected to in vivo and ex vivo IR. In general, experiments with

transgenic animals suggest a cardioprotective role for both Grx isoforms; however,

additional studies are needed to link the effects of each transgene to the protein

glutathionylation status of specific proteins and the functional consequences.

The first group to investigate the role of Grx on IR injury tested the effect of Grx1

embryonic knockout on infarct size and area at risk in an in vivo model of IR (Ho et al.,

2007). No difference in either parameter was observed in Grx1 knockout (KO) vs. WT animals, even though the de-glutathionylase activity of all of the mouse tissues, including

heart, was essentially absent. One possible explanation for this unexpected outcome is

that compensatory changes in the mechanisms of cellular homeostasis occurred during

development, offsetting the detrimental effect of Grx1 knock-out. Therefore, to

circumvent this complication, the Mieyal Laboratory recommends that future studies on

the effects of Grx1 in IR injury utilize a tissue-specific, inducible KO model.

Malik et al. (2008) further explored the role of Grx1 in IR injury by comparing the

effects of embryonic global glrx1 KO with muscle-specific overexpression, and by widening the scope of injury to include ischemic pre-conditioning (IPC) prior to IR. IPC is widely recognized to decrease subsequent IR injury (reviewed by Downey et al., 2007;

Hausenloy and Yellon, 2006), and there is evidence that Grx1 contributes to regulation of

several signaling pathways implicated in the mechanism of IPC (see below). Unlike Ho

et al. (2007), Malik and colleagues reported a small but significant increase in infarct size

(as well as decreased contractile performance) in glrx1 KO mice compared to controls.

In contrast to the effects of Grx1 KO, Grx1 overexpression appeared to decrease infarct

size and protect coronary function.

The basis for the distinct effects of glrx1 KO on infarct size reported by these two groups is not obvious, but likely reflects differences in experimental protocols. For example, Grx1 might play an important role in cardioprotection during early reperfusion, thus affecting infarct size after 2 hours of reperfusion (observed by Malik et al. (2008))

but exhibiting less of an effect after 4 hours (when Ho et al. (2007) measured infarct

size). Alternatively (or additionally), neurohumoral factors might have blunted the effect

of glrx1 KO in the in vivo IR model of Ho et al. (2007), while their absence revealed an

important cardioprotective role for Grx1 in the isolated heart model of Malik et al.

(2008). This scenario predicts that glrx1 KO indeed confers a detrimental effect on

cardiac function during IR, and compensation by other organ systems may represent an

important component of disease outcome when Grx1 activity is perturbed.

More dramatic than the role of Grx1 in IR injury was its contribution to IPC. Grx1

overexpression potentiated the protective effect of IPC on infarct size and cardiomyocyte

apoptosis, while IPC failed to confer any cardiac protection in Grx1 KO mice. To

explore the mechanism of Grx1-associated protection vs. IR injury, ROS production

(measured via malondialdehyde (MDA) content), Akt phosphorylation, and Bcl-2 content were assayed. Grx1 expression inversely correlated with MDA content following IR, and

Grx1 KO was associated with decreased IPC-induced Akt phosphorylation. The latter observation led the authors to suggest that Grx1 could contribute to IPC via activation of

Akt, which is consistent with other reports linking Grx activity to Akt phosphorylation

(Murata et al., 2003; Nagy et al., 2008; Wang et al., 2007), although the mechanism for this effect in vivo is not yet established (see Section 1.4.3, below). While Bcl-2 content was reported not to change with manipulation of Grx1, an apparent diminution of ~50% can be estimated by scrutiny of the bar graph which represents densitometric analysis of

Western blots from Grx1 KO mice (Malik et al., 2007; Figure 10).

Although Malik et al. (2007) observed an inverse correlation between Grx1 expression and MDA content, their conclusion that Grx1’s cardioprotective effects can be attributed to its role as an “antioxidant” is problematic. First, it is not consistent with the primary (and well-documented) function of Grx as a deglutathionylating enzyme

(Gravina and Mieyal, 1993; Chrestensen et al., 2000). In fact, studies of protein glutathionylation were not reported here, so it cannot be concluded that Grx1’s

cardioprotective effects were not related to its role in protein deglutathionylation.

Second, a Grx1-mediated antioxidant activity is not explicitly demonstrated. One could

imagine that, if Grx1 were expressed at high enough levels, it could serve as an

antioxidant by supplying reduced cysteines in a manner analogous to N-acetylcysteine or

DTT, but it is unlikely that such concentrations could be reproduced in vivo, even within

a disease context. Is there a more likely explanation for the decreased MDA content

observed in Grx1 transgenic mice subjected to IR? One possibility is that Grx1 may

indirectly decrease ROS by deglutathionylating a ROS-generating enzyme, such as complex I, which exhibits increased superoxide production upon glutathionylation

(Taylor et al., 2003). This interpretation is more consistent with documented activities of

Grx1, and could be investigated by determining the glutathionylation status of complex I during IR in WT and Grx1 transgenic mice. A similar scenario has been described for the cardioprotective role of TRx1 function in ischemic preconditioning. Analogous to glrx1 KO, inhibition of TRx1 resulted in increased MDA formation in the IR heart, suggesting an “antioxidant” function for the enzyme (Das, 2004). Instead of proposing

that TRx itself serves as an antioxidant, the authors suggested that it might promote

antioxidant activity via upregulation of antioxidant enzymes such as MnSOD. In light of

their distinct enzymatic activities, it is intriguing that Grx1 and TRx1 confer similar

cardioprotective effects during IPC, and determining the mechanism(s) for each

enzyme’s protective effects represents a fascinating avenue for future study.

The role of Grx2 in IR injury was also investigated using a transgenic animal model

(Nagy et al., 2008). As with Grx1 overexpression, Grx2 transgenic mice exhibited decreased infarct size and improved myocardial function following ex vivo IR compared

87 to WT mice. Unlike Grx1, Grx2 appeared to influence apoptosis, with Grx2 transgenic mice showing fewer apoptotic cardiomyocytes following IR compared to control animals.

Additional evidence supporting a cardioprotective effect of Grx2 included decreases in caspase activation, cardiolipin loss, MDA formation, and diminution of the GSH:GSSG ratio following IR. The roles of Grx2 in cytochrome c release and Akt phosphorylation were less straightforward. Grx2 transgenic animals exhibited decreased IR-induced cytochrome c release, but the amount of cytosolic cytochrome c was much higher at baseline in transgenic animals and appeared to decline with IR. Grx2 overexpression has been linked to decreased cytochrome c release in oxidant-challenged HeLa cells

(Enoksson et al., 2005), but the basis for its loss from the cytosol in Grx2 transgenic hearts with IR is puzzling. Phosphorylated Akt was also higher at baseline in transgenic vs. WT animals, with levels remaining steady following IR. Considering the distinct subcellular localizations of Grx2 and Akt (Grx2 in the mitochondrial matrix (Pai et al.,

2007) and Akt in the cytosol), this regulation is most likely indirect; however, a direct role of Grx2 in regulating Akt activity (as has been proposed for Grx1 (Murata et al.,

2003)) is possible if the Grx2 transgene was also expressed in cytosol. Since cytosolic

Grx activity was not reported in the study, this explanation cannot be ignored. In fact,

Enoksson and co-workers (2005) showed that Grx2 targeted to the cytoplasm protected

HeLa cells from doxorubicin-induced apoptosis, so verification of appropriate subcellular localization is a critical prerequisite for interpreting the findings of any study in which

Grx2 is overexpressed.

Overall, the work of Nagy et al. (2008) suggests a role for Grx2 in cardioprotection, but as in the case of other studies documenting the cytoprotective effect of Grx2 (Lillig et

88 al., 2004; Enoksson et al., 2005), mechanism(s) remain unknown. Importantly, candidate target pathways were proposed, including Akt and NFκB. Future work should focus on identifying the direct targets of Grx2 responsible for its pleiotropic effects on oxidant- induced signaling. An attractive candidate is mitochondrial complex I, in which glutathionylation of the 51- and 75-kD subunits is correlated with electron transport inhibition and increased production of superoxide (Taylor et al., 2003). The 51kD subunit contains one conserved cysteine that is not bound to an FeS cluster, and this cysteine faces the mitochondrial matrix (Hirst et al., 2003), making it a potential site of regulation by Grx2. Glutathionylation of complex I, with associated increases in superoxide production, would be expected to increase cytochrome c release and caspase activation, induce survival signals, and contribute to infarct size and cardiac dysfunction.

Thus, complex I deglutathionylation by Grx2 is a conceivable upstream event responsible for modulating these effects in Grx2 transgenic animals.

Finally, the roles of both Grx isoforms in IR injury were explored in rats by

Mukherjee et al. (2008). These investigators examined the effects of broccoli extract

(administered by oral gavage) on IR injury in isolated working rat hearts. Rats fed normal diets exhibited decreased Grx expression during IR, while rats given broccoli extract showed preservation of Grx1 and Grx2 content, which correlated with decreases in infarct size, cardiomyocyte apoptosis, cytochrome c release and caspase-3 activation, as well as improved post-MI cardiac and hemodynamic function. It is difficult, however, to determine the specific contribution of Grx isoforms to these cardioprotective effects, since broccoli gavage also induced other genes known to regulate cellular survival and

redox homeostasis (e.g., thioredoxins 1 and 2, thioredoxin reductase), and the effect of

broccoli gavage on protein glutathionylation status was not reported.

In summary, cardiac ischemia-reperfusion results primarily in increased protein glutathionylation; however, some proteins (e.g., complex II) are deglutathionylated during an IR episode. The effects of glutathionylation on function of these proteins (e.g.,

GAPDH, actin) may protect them from irreversible damage, or contribute to IR-induced injury. The latter possibility is supported by the cardioprotective phenotype of Grx transgenic animals, which exhibit decreased infarct size and improved cardiac function following IR. While data from KO animals are not as straightforward, studies on mice with inducible, tissue-specific KO might clarify the roles of Grx in acute IR injury. A critical frontier in this endeavor is to determine the mechanism(s) of Grx-related cardioprotection, which will require linking the protective effects of Grx on cardiac function with glutathionylation status of its molecular targets.

1.4.3 Preconditioning

Ischemic preconditioning (IPC) describes the phenomenon in which a series of brief ischemic episodes protects against subsequent IR injury (Murry et al., 1986;

Hausenloy and Yellon, 2006; Downey et al., 2007). Cardiac preconditioning can also be achieved by tachycardic stimuli, such as pacing or exercise (Domenech et al., 1998;

Sanchez et al., 2008). Potential targets of glutathionylation in preconditioning will be discussed below.

Sanchez et al. (2005; 2008) provide evidence that glutathionylation of the cardiac ryanodine receptor (i.e., RyR2) contributes to tachycardia- and exercise-induced preconditioning. Exercise and tachycardic pacing both result in RyR2 glutathionylation,

90 which is correlated with increased Ca2+ release rates (Sanchez et al., 2005) and increased colocalization with NADPH oxidase (Sanchez et al., 2008). NADPH oxidase inhibitors prevent these effects, suggesting that ROS production by this enzyme is the likely trigger for RyR2 glutathionylation in vivo. Importantly, NADPH oxidase inhibitors also abolish the protective effect of tachycardia or exercise on infarct size. While RyR2 glutathionylation is not explicitly correlated with these observed changes, they are consistent with the concept that RyR2 glutathionylation contributes to tachycardic or exercise-induced preconditioning. This concept could be tested further by transfection of a mutant RyR2 that cannot be glutathionylated and determining the effect on tachycardic

PC, or determining the role of Grx1 on RyR2 glutathionylation status (assuming the site of glutathionylation is exposed to the cytosol).

Unlike tachycardic or exercise-induced PC, a specific role for protein glutathionylation has not been established for IPC. However, the apparent contribution of Grx1 to IPC (via TG overexpression or KO, see Malik et al. (2008) and above) implicates glutathionylation as a potential modulating mechanism. Numerous signaling intermediates have been implicated in the regulation of IPC, of which many components have been shown in other systems (reviewed by Shelton et al. (2005) to be regulated by

S-glutathionylation; hence, modulation of these pathways by glutathionylation in the heart represents a potential layer of regulation for cardiac IPC.

1.4.3A. Protein Kinase C (PKC) –

PKC is considered to be a central regulator of IPC signaling (Downey et al.,

2007), integrating multiple upstream signals (e.g., catecholamines, angiotensin II, endothelin) and activating diverse downstream pathways implicated in IPC reviewed by

(reviewed by Hausenloy and Yellon, 2006). Importantly, most PKC isoforms appear to be sensitive to inhibition by S-glutathionylation (reviewed in (Shelton et al., 2005), including those isoforms implicated in IPC signaling (Hausenloy and Yellon, 2006).

Therefore, a potential mechanism by which Grx1 contributes to IPC may be via deglutathionylation (and subsequent activation) of PKC. This pathway could be tested by determining the effect of Grx manipulation on PKC activity, and correlating these changes to its glutathionylation status during brief ischemic challenges.

1.4.3B. Protein Kinase A (PKA) –

Sanada et al. (2004) provided evidence that PKA may contribute to IPC independently from PKC, presumably through inhibitory effects on Rho kinase and cytoskeletal reorganization. PKA is inhibited by glutathionylation in vitro, as well as in mouse fibroblasts treated with diamide (Humphries et al., 2002; reviewed in Shelton et al., 2005). As for PKC, Grx could contribute to IPC via maintaining PKA in its active, deglutathionylated form. However, the effects of IPC on PKA glutathionylation status

(and the additional role of Grx) have not yet been explored.

1.4.3C. Nuclear Factor κB (NFκB) –

NFκB is a pleiotropic transcription factor commonly activated during cellular stress. While the effects of NFκB activation are often dependent upon cell type and stress stimulus, its activation has been shown primarily to increase survival in cardiomyocytes

(Jones et al., 2003). Two studies by Maulik and colleagues (Maulik et al., 1998; Maulik et al., 1999) utilizing an inhibitor of NFκB nuclear translocation suggest that the pathway is required for IPC in the isolated, perfused rat heart, and that the basis for its protective effect may be through activation of Bcl-2 transcription. Various groups have provided

evidence that the NFκB pathway is regulated at various foci by glutathionylation (e.g., p50 (Pineda-Molina et al., 2001), p65 (Qanungo et al., 2007), IKK (Reynaert et al., 2006;

Shelton et al., 2009), and the proteasome (Obin et al., 1998)), with glutathionylation having an inhibitory effect in each case. Therefore, deglutathionylation of p50, p65, IKK, or the proteasome by Grx could maintain overall pathway activity during IPC. As in the examples discussed above, establishing such a role for Grx will require analysis of glutathionylation status of NFκB pathway components during IPC, and correlating changes in glutathionylation status with corresponding changes in activity.

1.4.3D. Akt –

Akt, also known as PKB, has been implicated in cell growth, survival, and migration signals in diverse cell types. Numerous studies have identified Akt activation as contributing to both early (i.e., PKC activation) and late (i.e., mitochondrial) events in

IPC (Hausenloy and Yellon, 2006). While Akt itself has not been shown to be glutathionylated, it appears to be regulated (either directly or indirectly) by Grx1. Murata et al. (2003) reported that H9c2 cells (rat embryonic cardiomyocytes) in which Grx1 was overexpressed exhibited decreased H2O2-induced apoptosis, and this effect was correlated

with an increased duration of Akt phosphorylation and decreased association with PP2A,

the phosphatase associated with Akt inactivation. Studies on isolated Akt in the presence of H2O2, Grx, and/or GSH were interpreted to mean that Grx directly reduces an Akt intramolecular disulfide in vivo; however, the likelihood of this representing a physiological regulatory mechanism is questionable, since in vitro conditions did not approximate the cellular environment (e.g., GSH:GSSG = 20, lower than the typical ratio), kinetic competence was not demonstrated (endpoint assay was 30 min.), turnover

conditions were not used ([Grx1] was 80-fold higher than Akt), and Akt redox status was

shown not to affect its kinase activity. Furthermore, as described above, the specificity of

Grx for glutathionyl mixed disulfides would preclude its action on an intramolecular

disulfide substrate. A more likely mechanism of Grx-mediated regulation of Akt is via

Akt’s phosphorylation status, as has been suggested by other groups exploring the effect

of Grx on Akt activity and post-translational modification. For example, Akt

phosphorylation is increased in Grx2 transgenic mice (Nagy et al., 2008), and Grx1 overexpression is correlated to increased Akt phosphorylation in bovine aortic endothelial cells (Wang et al., 2007). The mechanism of regulation of Akt phosphorylation by Grx is still not resolved. However, the latter authors appropriately consider that the level of regulation of Akt could be direct (as proposed by Murata’s group) or indirect, via deglutathionylation of upstream activators such as PKA or PKC.

In summary, although a role for reversible glutathionylation in IPC has not yet been established, it is suggested by the contributory role of Grx1 in a transgenic animal model (Malik et al., 2008). Here, signaling candidates were identified and discussed which are implicated in IPC and also are established targets of regulation by S- glutathionylation, including PKC, PKA, and NFκB pathway components, all of which are rendered inactive upon glutathionylation. Akt was also discussed as being regulated by

Grx, although the specific nature of the regulation is not yet understood. Importantly, no study to date has reported global or specific protein glutathionylation during IPC. Future work should focus on potential links between glutathionylation status of these proteins, and the effects of glutathionylation on their functions as well as on IPC in general.

A final consideration in the discussion of ischemic vs. tachycardic PC is the

potential roles of Grx. Although the mechanism remains unknown, Grx1 appears to

contribute to the protective effects of IPC (Malik et al., 2008); however, one might

predict that Grx would hinder tachycardic PC via deglutathionylation of RyR2.

Importantly, the effect of Grx1 activity on RyR2 glutathionylation status has not been reported, contrasting with documented Grx-dependent de-glutathionylation of RyR1

(Aracena-Parks et al., 2006). Determining the effect of manipulated Grx levels on RyR2 glutathionylation status (and Ca2+ release rates) would help address this question.

1.4.4 Nonspecific oxidative injury

IR and IPC are both conditions of oxidative stress. In IR, ROS are produced primarily from Complexes I and III of the mitochondrial electron transport chain

(Lesnefsky et al., 2004; Lesnefsky and Hoppel, 2003), and ROS generation from mitochondria and/or NADPH oxidase appears to contribute to PC signaling (Downey et

al., 2007; Sanchez et al., 2008). Above, we discussed evidence for increased

glutathionylation of the intracellular proteome—as well as individual proteins—with IR, and the potential for regulation of survival proteins by glutathionylation during IPC.

Importantly, additional proteins are reported to be regulated by glutathionylation during

generalized oxidative challenges, such as H2O2 treatment or exposure to decreased

GSH:GSSG ratios. Although these oxidative stimuli do not necessarily model the

physiological state or a specific disease condition, they identify candidate proteins for

regulation by glutathionylation during pathological oxidative stresses such as IR, as well

as other cardiovascular diseases associated with oxidative stress, such as hypertension

(Harrison et al., 2007) and atherosclerosis (Kher and Marsh, 2004).

For example, mitochondrial complex I is glutathionylated in vitro upon exposure to GSSG (Taylor et al., 2003) or low GSH:GSSG ratios (i.e., 0.67-12, (Beer et al.,

2004)), and glutathionylation is reversed upon incubation with Grx2 and GSH. Complex

I glutathionylation results in increased superoxide production (Taylor et al., 2003), suggesting that this modification would increase ROS generation, leading to activation of redox signaling pathways and/or induction of cell death, depending upon the magnitude of modification. Key considerations regarding the possibility of complex I regulation by glutathionylation in vivo include its mechanism of formation and its potential role in cardiac disease.

Taylor et al. (2003) propose that oxidative stress within the mitochondria alters the GSH:GSSG ratio sufficiently to cause complex I-SSG formation by thiol disulfide exchange; however, this mechanism is unlikely unless the modified cysteines display unusually low redox potentials (Gallogly and Mieyal, 2007; Mieyal et al., 1995; Gilbert,

1995). Thus, alternative mechanisms of glutathionylation (e.g., via nitrosothiol intermediate, sulfenic acid intermediate, etc., as described above) are more probable. An additional mechanism of glutathionylation was proposed by Beer et al. (2004), namely, catalysis by Grx2. Although mammalian glutaredoxins are efficient protein deglutathionylating enzymes, Grx1 promotes protein glutathionylation in the presence of

GS∙ radical (Starke et al., 2003) and—to a much lesser extent—GSNO or GSSG. To determine if Grx2 exhibited similar behavior, it was incubated with 5 mM GSSG and mitochondrial membranes from rat heart containing complex I. Addition of Grx2 accelerated glutathionylation of membrane thiols over a short time course; however, when GSH was added to glutathionylated membrane proteins, Grx2 incubation led to

96 overall deglutathionylation of protein-SSG (Beer et al., 2004). Since the latter conditions more closely represent the intermitochondrial milieu, they better reflect the potential environment in which Grx2 may regulate complex I-SSG in vivo. Thus, catalysis of glutathionylation by Grx2 involving GSSG appears not to be a likely mechanism of complex I-SSG formation. However, Grx2-mediated GS-radical transfer may be a more feasible mechanism (see Chapter 3).

Whether complex I is indeed regulated by glutathionylation in the intact heart has not yet been explored. Studies focused on documenting complex I-SSG formation in cardiac cells or tissue, with an oxidative stimulus relevant to cardiac disease (e.g., IR, angiotensin II treatment), and attention to the effects of Grx1 and Grx2 on complex I glutathionylation status will provide additional insight into this potential contribution to cardiac injury.

Another mitochondrial enzyme potentially regulated by glutathionylation during cardiac oxidative stress is α-ketoglutarate dehydrogenase (KGDH). Nulton-Persson and colleagues (2003) demonstrated that H2O2 treatment of rat heart mitochondria led to inhibition of KGDH activity, which was reversed by Grx1 and GSH within minutes, but unaffected by the TRx system. Although KGDH glutathionylation was not shown directly, it was inferred from the recovery of activity by Grx1 treatment, and hypothesized to protect catalytic cysteine residues from irreversible damage during oxidative stress conditions, such as IR.

A later study by the same group (Applegate et al., 2008) focused on the glutathionylation site of KGDH, and proposed an intriguing model in which glutathionylation occurs on a covalently bound lipoic acid moiety, rather than the typical

protein-cysteine sulfhydryl moiety. This conclusion was based on observations that H2O2

treatment prevented recognition of KGDH by an anti-lipoate antibody, as well as HNE-

mediated oxidation. A key question concerning the proposal of mixed disulfide

formation between KGDH-lipoic acid and GSH is its mechanism of stabilization. It

would be expected that the vicinal thiol on lipoic acid would undergo thiol-disulfide exchange with its neighboring R-SSG moiety, forming lipoic acid intramolecular disulfide and GSH. Structural analysis or modeling studies might identify potential residues that stabilize the second, reduced thiol on lipoic acid, making it unavailable to react with the neighboring R-SSG. While stable mixed disulfide formation between protein-bound lipoic acid and GSH is a novel concept, and catalysis of lipoic acid-SSG would represent a new activity for Grx1, there are alternative interpretations to the authors’ observations. For example, it is possible that KGDH is glutathionylated on a cysteine residue in close proximity to the bound lipoic acid, and steric interference by this glutathionylated cysteine blocks accessibility of lipoic acid to antibodies and HNE.

Alternatively, glutathionylation on a distant Cys could induce a conformational change with the same effect on access of the lipoic acid to detection reagents. This possibility could be addressed by analyzing the glutathionylated product by mass spectrometry, and/or by isolating the lipoyl moiety prior to analysis for S-glutathionylation.

Overall, glutathionylation of complex I and KGDH appear to be facile upon exposure of mitochondria to oxidants in vitro; however, their relevance to oxidative stress-associated cardiac disease is not yet established. Determination of their glutathionylation status with IR, or other pathophysiological oxidative challenge, will provide insight into the likelihood of their regulation by glutathionylation in vivo.

1.4.5 Cardiac Hypertrophy

As for IPC, multiple signaling pathways contribute to the development of

pathological cardiac hypertrophy (reviewed by Heineke and Molkentin, 2006). Among

them is the Raf/MEK/ERK pathway, which can be stimulated either by G protein-

coupled receptor (GPCR) ligands (e.g., angiotensin II, endothelin) or by mechanical

stretch, resulting in induction of protein synthesis.

Recently, Pimentel et al. (2006) showed that mechanical strain-stimulated

Raf/MEK/ERK pathway activation in neonatal rat ventricular myocytes was dependent

upon glutathionylation (and subsequent activation) of Ras, a small GTPase implicated in

myocyte growth signaling (see Figure 1.6, p. 135, right). Notably, this study is one of

few that fulfill most of the criteria for establishing S-glutathionylation as a regulatory mechanism. Specifically, the authors demonstrated that Ras-SSG formed in response to a physiological stimulus (mechanical strain) under physiological conditions (intact cells assumed to have a normal GSH:GSSG ratio); that Ras glutathionylation conferred effects on protein function (increased Raf and GTP binding); that Ras-SSG formation was reversed by Grx (documented via overexpression); and that Ras glutathionylation impacted a downstream pathway important in disease (protein synthesis in cardiac hypertrophy). The proposed role of Ras-SSG in hypertrophic signaling is summarized in

Figure 1.6, p. 135.

Determination of Ras glutathionylation status in animal models of cardiac hypertrophy would provide insight into the relevance of this pathway in progression of the disease in vivo. An animal model would also allow exploration of the effects of

99 chronic strain—vs. acute strain—on Ras-SSG glutathionylation status, activity, and cardiac hypertrophy.

1.4.6. Atherosclerosis

Atherosclerosis is a complex disease process involving interactions between multiple cell types in the blood and vasculature. The precise role of glutathionylation in the development and progression of atherosclerosis is unknown; however, conditions within atherosclerotic plaques (e.g., hypoxia, oxidative stress, oxLDL, and inflammation) have been shown in other contexts to promote glutathionylation (Qanungo et al., 2007;

Eaton et al., 2002; Wang et al., 2006; Sullivan et al., 2000), and Grx has been reported to associate with areas of oxidative stress within the vasculature (Okuda et al., 2001). The following discussion explores further evidence for involvement of protein glutathionylation in atherogenesis.

Global protein glutathionylation increases in human monocyte-derived macrophages exposed to oxidized LDL (oxLDL) (Wang et al., 2006), a major component of atherosclerotic plaques also believed to contribute to their progression (Kher and

Marsh, 2004). Together with GSH depletion, increased protein-SSG content was implicated in oxLDL-induced macrophage death in vitro. Dying macrophages represent a major component of atherosclerotic plaques, and their presence in atherosclerotic lesions increases risk of rupture (Lee and Libby, 1997). The role(s) of specific glutathionylated proteins in macrophage cell death is not yet determined, nor is it known whether global protein glutathionylation increases in other cells types exposed to oxLDL; however, these questions form the basis for future studies.

100

Nonaka et al. (2007) discovered that patients with arteriosclerosis of the

extremities (i.e., arteriosclerosis obliterans, ASO), exhibit increased glutathionylation of

serum proteins detected by SDS-PAGE followed by GST overlay. Remarkably, there was a positive correlation between disease progression and magnitude of protein glutathionylation measured, leading the authors to conclude that serum protein glutathionylation is both a sensitive and specific marker of ASO. However, many of the patients enrolled in the study had comorbid conditions also associated with increased serum protein-SSG, such as tobacco use (Muscat et al., 2004), making the specificity of this marker for ASO unlikely. Importantly, these authors identified the serum protein apoB100 as a target for increased glutathionylation in ASO. ApoB100 is the major component of LDL, and it is tempting to speculate that its glutathionylation could affect its function, as has been shown for other post-translational modifications (Swift, 1996).

Whether apoB100-SSG simply represents a disease marker, or contributes to the pathogenesis of ASO, remains an open question.

In the case of SERCA, the sarcoplasmic reticulum calcium ATPase, glutathionylation appears to be part of a normal regulatory mechanism that is disrupted during atherosclerosis. Adachi and colleagues (2004b) demonstrated that SERCA glutathionylation occurs in vascular cell lines and tissues in the presence of RNS and endogenous GSH. Moreover, glutathionylation could be stimulated by physiological ligands known to generate RNS (e.g., acetylcholine, bradykinin). SERCA glutathionylation increased its ATPase activity, correlated with vessel dilation, and occurred in the presence of a cGMP inhibitor, leading the authors to propose that SERCA glutathionylation represents a physiological, cGMP-independent mechanism of vessel

101

relaxation. Site-directed mutagenesis and mass spectroscopic analysis suggested that

glutathionylation of Cys674, located in the cytosolic-facing hinge domain, was

responsible for SERCA activation. Interestingly, analysis of cysteine modifications from

atherosclerotic vs. normal rabbit aortas indicated increased sulfonate formation (including

C674), which corresponded to decreased NO-induced relaxation, glutathionylation, and

Ca2+ reuptake. Taken together, these observations suggest that irreversible oxidation

(i.e., sulfonic acid formation) of SERCA’s C674 during atherosclerosis prevents

regulation of function by reversible glutathionylation and may contribute to the impaired

vasodilation response to NO in atherosclerotic smooth muscle.

Adachi et al. (2004a) demonstrated that glutathionylation of Ras may contribute

to vascular hypertrophy (implicated in atherosclerosis and hypertension) by activating

protein synthesis in rat vascular smooth muscle cells (VSMCs). Treatment of VSMCs

with angiotensin II (AII), an established stimulus for vascular hypertrophy, led to

glutathionylation and activation of Ras, which resulted in increased phosphorylation of

p38 and Akt, and increased protein synthesis (Figure 1.6, p. 135, left). These effects

were dependent upon NADPH oxidase activation and ROS formation (shown separately

to be activated by AII (Wang et al., 2001a; Landmesser et al., 2002)), and were blocked

by overexpression of Grx1 or mutation of Ras at the site of glutathionylation (C118).

Interestingly, AII-stimulated ERK activation, which contributes to AII-induced protein

synthesis, was not redox-sensitive and proposed to occur independently of Ras

glutathionylation. Like the work of Pimentel et al. (2006), the work of Adachi and

colleagues represents an excellent demonstration of protein regulation by S- glutathionylation: Ras-SSG formed in response to a physiological stimulus (AII

102

treatment); glutathionylation resulted in a change in protein activity (increased Raf

binding); was reversed by Grx1 (documented via overexpression); and was correlated to a

physiological outcome (protein synthesis).

The work of Adachi and Pimentel point to several common events in hypertrophic

signaling within the heart and vasculature: both require production of endogenous H2O2,

result in Ras glutathionylation, and activate signaling pathways that ultimately result in

increased protein synthesis. However, it is intriguing that the Ras-SSG-activated

pathways implicated in hypertrophy differ in the two model systems. Why might H2O2- induced Ras glutathionylation activate the Raf/MEK/ERK pathway in cardiomyocytes vs. p38 and Akt—but not ERK—in vascular smooth muscle? (Figure 1.6, p. 135) The

answer could reflect differences in signal transduction networking between cell types,

different degrees of Ras glutathionylation resulting from each stimulus (assuming

different thresholds of activation for downstream pathways), and/or distinct localization of ROS production (and subsequent Ras glutathionylation) depending upon the nature of the stimulus (i.e., NADPH oxidase vs. the source of strain-stimulated ROS).

In addition to modulating AII signaling in VSMCs, Ras-SSG may contribute to atherosclerosis by mediating the response to oxLDL in endothelial cells (see Figure 1.7, p. 136). Clavreul et al. (2006a) demonstrated that treatment of bovine aortic endothelial cells (BAECs) with peroxynitrite led to Ras glutathionylation and activation of both ERK and Akt pathways, and some of these observations were recapitulated with oxLDL treatment. Unlike mechanical strain- and AII-induced Ras glutathionylation, which required formation of H2O2, oxLDL-mediated glutathionylation was dependent upon

peroxynitrite. The authors argue for a thiol-disulfide exchange mechanism with GSSG,

103

based on proposed chemical reactions between peroxynitrite and GSH (producing

GSSG), which exhibit a time course of GSSG formation compatible with that observed

for Ras glutathionylation. This mechanism is feasible if the Kmix for Ras-Cys118-SSG

formation is similar to the GSH:GSSG ratio achieved during peroxynitrite treatment

(Gilbert, 1990; Gilbert, 1995); but GSSG concentration was not measured in this study,

and to the best of our knowledge, the Kmix for Ras-C118 has not been determined.

Therefore, the mechanism of Ras glutathionylation associated with peroxynitrite

treatment remains unclear.

Specific physiological effects of Ras glutathionylation (and subsequent activation of MEK/ERK and Akt) that might contribute to atherosclerosis were not reported here; however, it has been reported in other contexts that MEK/ERK activation within the endothelium may induce proliferation of endothelial cells, contributing to atherogenic

vascular remodeling (Pintus et al., 2003).

Although a mechanism for endothelial Ras-SSG-induced atherogenesis was not fully elucidated by Clavreul et al. (2006a), a role for oxLDL-induced Ras glutathionylation in insulin resistance was described in a subsequent study by the same group (Clavreul et al., 2006b). Here, the effects of oxLDL-induced Ras glutathionylation were followed over a longer time course, and cross-talk with a second signaling pathway

(insulin/insulin-receptor substrate (IRS)/Akt) was explored. Here, as in the previous study (Clavreul et al., 2006a), oxLDL-induced Akt activation was transient, peaking at 15 minutes; however, ERK activation was sustained (>1 hour). Moreover, subsequent activation of Akt by insulin/IRS was blunted by pretreatment with oxLDL, presumably via ERK-mediated phosphorylation (and inactivation) of IRS (Figure 1.7, p. 136, right).

104

This work enhances understanding of downstream effects of Ras-SSG in endothelial cells, particularly within the context of insulin resistance. However, the distinctive effects of peroxynitrite and oxLDL on the time course of ERK activation raise questions about the use of exogenous peroxynitrite to represent a physiologically relevant stimulus.

While studied in different vascular cells using different oxidative stimuli, both

Adachi and Clavreul report Ras-SSG-mediated phosphorylation of Akt, which is diminished by overexpression of Grx1. Interestingly, Wang et al. (2007) report an opposite correlation between Grx1 activity and Akt activation in BAECs exposed to laminar flow. Grx1 activity approximately doubled within 5 minutes of exposure to physiological flow rates, and this activation correlated to increased phosphorylation of

Akt and eNOS. Akt and eNOS phosphorylation were augmented with overexpression of

Grx1, and diminished after treatment with Grx1 siRNA, suggesting that Grx1 activity regulates their activation, although a specific mechanism was not identified. These observations are consistent with those of Murata et al. (2003), who reported increased

Akt phosphorylation in H9c2 cardiomyoblasts overexpressing Grx1. Taken together, these studies highlight the complex relationship among Grx activity, protein glutathionylation, and Akt activity within the cardiovascular system. Importantly, Akt is emerging as a complicated signaling molecule within the heart and vasculature, implicated in various pathological signaling events as well as in normal development and homeostasis (Mullonkal and Toledo-Pereyra, 2007). It is conceivable that Grx could participate in regulating the balance between physiological (i.e., laminar flow-induced) and pathophysiological (i.e., ATII-induced) Akt activation. Determining the status of

105

Akt activation (as well as downstream effects such as eNOS activity and vessel

hypertrophy) in Grx TG and KO animals would help address this complex situation.

An emerging contributor to atherogenesis is tumor necrosis factor-alpha (TNFα), which is thought to induce expression of adhesion molecules on endothelial cells and contribute to vascular smooth muscle cell apoptosis (Dixon and Symmons, 2007). Pan and Berk (Pan and Berk, 2007) treated endothelial cells with a combination of TNFα and cycloheximide (CHX), and observed Grx activation, pro-caspase-3 deglutathionylation, caspase-3 cleavage and increased apoptosis. These effects were blocked by transfection with siRNA against Grx1, leading the authors to propose that Grx1-mediated deglutathionylation of pro-caspase-3-SSG contributes to TNFα-induced apoptosis.

Importantly, this study was the first to demonstrate glutathionylation of pro-caspase-3 and its effect on susceptibility to cleavage, and this report also identifies caspase-3 as another protein that exists in a glutathionylated state under resting conditions, becoming deglutathionylated by Grx1 in response to an ROS-generating stimulus.

Still, there are some difficulties in interpreting the results of this work. First, are the effects on caspase-3 glutathionylation due to TNFα or CHX? This is difficult to answer because the treatment given to control cells is not explicitly stated. Second, how closely does the TNFα concentration given to BAECs compare to circulating levels in diseased vessels? Here, a dose response curve of response to TNFα would be informative. Finally, this study raises an important question about the potential role of

Grx in atheroprotection. In the case of Ras, Grx levels were correlated with decreased

Ras-SSG and decreased hypertrophic signaling. In the case of pro-caspase-3, however,

Grx overexpression (and deglutathionylation) led to increased apoptosis, a presumably

106

atherogenic event. Taken together, these results highlight the fact that the role of Grx in

cardiovascular disease may not be entirely straightforward, with its roles in disease

protection or progression dependent upon cell type, extracellular stimuli, etc.

1.5 REGULATION OF APOPTOSIS BY GLUTATHIONYLATION AND

GLUTAREDOXINS

Several recent studies focused on the roles of Grx in apoptotic regulation (see Table

1.5, p. 124) indicate that both Grx isoforms regulate apoptosis under particular conditions in different cell types. Grx1 either sensitizes or protects cells from apoptosis, depending upon the circumstances. In the context of H2O2 treatment, both overexpression and inhibiton studies indicate that Grx1 is protective (Murata et al., 2003; Chrestensen et al.,

2000). Work by Murata’s group suggests that the protective mechanism may involve

regulation of Akt activity, possibly via deglutathionylation (Mieyal et al., 2008).

Deglutathionylation of another apoptotic mediator (procaspase-3) is implicated in the

sensitizing effect of Grx1 towards TNFα-induced apoptosis in fibroblasts (Ho et al.,

2007) and endothelial cells (Pan and Berk, 2007), while caspase 3-catalyzed cleavage of

Grx1 appears to sensitize lung epithelial cells to Fas-induced apoptosis (Anathy et al.,

2009). It appears from these studies that Grx1 may exhibit opposing effects on apoptotic

susceptibility depending on the model system and apoptotic stimulus. For Grx2, two

studies to date suggest anti-apoptotic functions for the enzyme (Lillig et al., 2004;

Enoksson et al., 2005), although both were performed in the same cell line and utilized some of the same oxidative stimuli. The mechanism(s) underlying Grx2’s cytoprotective

effect seem to involve regulation of mitochondrial integrity, but the target of regulation

and the catalytic activity required are unknown.

107

1.6 BCL-2 AND BCL-XL

Bcl-2 and Bcl-xL represent two anti-apoptotic members of the Bcl-2 protein family.

They are structurally similar, containing four Bcl-2 homology domains and a transmembrane domain. Bcl-2 and Bcl-xL are implicated in protection against apoptosis triggered by diverse stimuli in multiple cell types, including cardiomycoytes (reviewed in

(Kim, 2005; Gustafsson and Gottlieb, 2007; Gross et al., 1999). Early studies in T cells suggested that Bcl-2 and Bcl-xL exhibited redundant roles, and it was concluded that they represent essentially interchangeable anti-apoptotic factors (Chao et al., 1995). However,

recent discoveries indicate that Bcl-2 and Bcl-xL exhibit only partially overlapping roles in cytoprotection (see Table 1.6, p. 126), with Bcl-xL potentially acting as a more potent anti-apoptotic molecule compared to Bcl-2 (Fiebig et al., 2006). Additional support for distinct roles of Bcl-2 and Bcl-xL in cardiomyocyte apoptosis will be presented in

Chapter 4.

108

Scheme 1.1. Potential spontaneous chemical reactions leading to protein glutathionylation. Evidence supporting each reaction pathway is presented in Section

1.1. Protein-SSG, protein-glutathione mixed disulfide.

109

Scheme 1.2. Reactions of mammalian glutaredoxins. A, Catalytic mechanism of

deglutathionylation by human glutaredoxins (Gravina and Mieyal, 1993; Srinivasan et

al., 1997; Gallogly et al., 2008). In the first step, the thiolate of Grx’s N-terminal active

site cysteine attacks the glutathionyl sulfur of the protein-glutathione mixed disulfide (P-

SSG), forming the Grx-SSG intermediate and releasing reduced protein-SH (P-SH). In

the second step, free GSH attacks the glutathionyl sulfur of the Grx-SSG intermediate,

releasing reduced Grx and GSSG. GSSG is then reduced to 2GSH by GSSG reductase

(GR) and NADPH. Step 3 represents a side reaction in which Grx’s C-terminal active

site cysteine competes with GSH for reduction of Grx-SSG, forming a Grx active site disulfide and releasing GSH. The Grx-S2 side-product is reduced by GSH and recruited

back into the catalytic cycle. B, Grx can utilize GSSG as an oxidized substrate (Gallogly

et al., 2008; Starke et al., 2003; Mieyal et al., 1991b). This scheme is analogous to

Scheme 1A, except that the first substrate for Grx is glutathionylated glutathione (i.e.,

GSSG), and the second substrate is protein-SH. This reaction occurs under oxidizing

conditions (i.e., low GSH:GSSG ratio) until the protein-SH:protein-SSG ratio reaches equilibrium. C, Proposed mechanism of glutathione thiyl radical (GS•) scavenging by

Grx (Starke et al., 2003). In the first step, the N-terminal active site cysteine of Grx

attacks GS•, forming a Grx disulfide anion radical intermediate. This radical then reacts

•- with O2 in step 2, forming superoxide (O2 ) and the typical Grx-SSG intermediate. In step 3, the Grx-SSG intermediate is reduced by GSH, forming GSSG and reduced enzyme. D, Proposed mechanism of glutathionyl transfer by Grx (Starke et al., 2003). In

the first step of the reaction, the Grx catalytic cysteine thiolate attacks GS●, forming the

Grx-SSG●- disulfide anion radical intermediate. This intermediate can proceed to react

110 with protein-SH (P-SH, step 2), forming protein thiyl radical (P-S●). In step 3, another

Grx-SSG●- molecule reacts with P-S● (step 3), quenching the radical reaction and forming protein-SSG (P-SSG). The net reaction yields protein-SSG from 2 GS● and 2 P-SH molecules.

111

112

Scheme 1.3 Proposed mechanism for Prx glutathionylation by GSTπ (Manevich et

al., 2004). Prx sulfenic acid (Prx-SOH) is generated by reduction of phospholipid hydroperoxide (PL-OOH) to PL-OH. GSTπ mediates nucleophilic displacement of the

Prx-OH hydroxyl group by GSH, forming Prx-SSG and H2O. Prx-SSG is then reduced to

Prx-SH by GSH.

113

Scheme 1.4. Complete commitment to catalysis vs. encounter-type catalytic

mechanisms. A, In the case of high commitment to catalysis, there is a reversible

binding step between enzyme and substrate, followed by a chemistry step (k2) that is very

fast compared to the rate of enzyme-substrate dissociation (k-1). Thus, essentially every

substrate molecule that binds to enzyme undergoes a nucleophilic displacement reaction.

B, In the case of an encounter-type mechanism, enzyme and substrate react upon association, but without formation of a reversible complex. This latter model is supported by two-substrate kinetic analysis of Grx1 (Srinivasan et al., 1997) and Grx2

int (Gallogly et al., 2008), which predicts “true” KM values approaching infinity for both

substrates.

114

Table 1.1. Candidate enzymes for catalysis of protein S-glutathionylation.

Proposed role in Typical Reaction reversible Enzyme Catalyzed Reference glutathionylation Reference (Rodriguez- Manzaneque deglutathionylation et al., 1999; Grx5 uncertain N/A Shenton et (mechanism uncertain) al., 2002; Tamarit et al., 2003) PRx-SO2H + ATP + 2RSH  deglutathionylation (Findlay et (Jeong et SRx al., 2006; PRx-SH + ADP + al., 2006) (mechanism uncertain) Tew, 2007) R-SS-R + H2O ED + GSH  ED-SG (Manevich glutathionylation et al., 2004; (Hayes et GSTπ (ED = electron Ralat et al., al., 2005) (see Scheme 2B) deficient 2006; Tew, 2007) compound) glutathionylation:

E-(SH)2 + O2  E- E-(SH)2 + O2  S + H O E-S + H O 2 2 2 (Thorpe et 2 2 2 Section QSOX 2R-SH + E-S2  al., 2002) Protein-SH + GSH + E- 1.1.5C

R-SS-R + E-(SH)2 S2  Protein-SSG + E-

(SH)2 glutathionylation:

GPx-SeH + H2O2

GPx-SeOH + H2O H O + 2GSH  GPx-SeOH + GSH  GPx- 2 2 (Flohe, Section + like GSSG + H2O 1985) GPx-SeSG + H 1.1.5D GPx-SeSG + Protein-SH  Protein-SSG + GPx- SeH

115

Proposed role in Typical Reaction reversible Enzyme Catalyzed Reference glutathionylation Reference glutathionylation:

Protein-SH + O2  mono- RSH + O  R- (Abraham 2 Section oxygen- et al., Protein-SOH + H2O SOH + H2O 1.1.5D ase-like 1983) Protein-SOH + GSH 

Protein-SSG + H2O

116

Table 1.2. Deglutathionylation activities and glutathionyl specificity of

glutaredoxins from prototype organisms.

Gluta- CXX*C/S Deglutathionylation Glutathionyl redoxin motif activity? specificity?† Reference E. coli Grx1 CPYC Yes Y (Bushweller et (peptide-SSG, (Step 1) al., 1992; ArsC-SSG) Holmgren, 1976; Peltoniemi et al., 2006a; Shi et al., 1999) Grx2 CPYC Yes Y (Aslund et al., (βME-SSG) (Step 1) 1994; Vlamis- Gardikas et al., 1997) Grx3 CPYC Yes Y (Aslund et al., (βME-SSG) (Step 1) 1994) Grx4 CGFS No N/A (Fernandes et al., 2005)

S. cerevisiae Grx1 CPYC Yes ND (Gan, 1992; (βME-SSG) Luikenhuis et al., 1998; Discola et al., 2008)

Grx2 CPYC Yes ND (Discola et al., (βME-SSG, 2008; Silva et 20S proteasome- al., 2008; SSG) Luikenhuis et al., 1998) Grx3 CGFS Yes* ND (Rodriguez- Manzaneque et al., 1999)

117

Gluta- CXX*C/S Deglutathionylation Glutathionyl redoxin motif activity? specificity?† Reference Grx4 CGFS Yes* ND (Rodriguez- Manzaneque et al., 1999) Grx5 CGFS unresolved** ND (Rodriguez- Manzaneque et al., 1999; Rodriguez- Manzaneque et al., 2002; Tamarit et al., 2003; Shenton et al., 2002) Grx6 CSYS Yes Y (Mesecke et al., (Step 1) 2008) Grx7 CPYS Yes Y (Mesecke et al., (Step 1) 2008)

H. sapiens Grx1 CPYC Yes Y (Srinivasan et al., (protein-SSG, (Steps 1 & 2) 1997; Yang et al., cysteine-SSG, βME- 1998; Johansson SSG) et al., 2004; Gravina and Mieyal, 1993) Grx2 CSYC Yes Y (Johansson et al., (protein-SSG, (Steps 1 & 2) 2004; Gallogly et cysteine-SSG, βME- al., 2008) SSG) Grx5 CGFS ND ND (Molina-Navarro et al., 2006; Camaschella et al., 2007) Grx domains TGR CPHS Y Y*** (Sun et al., 2001; (βME-SSG) Sun et al., 2005) PICOT CGFS ND ND (Isakov et al., 2000; Witte et al., 2000) TR3 CTRC N N/A (Su and Gladyshev, 2004)

118

†Indicated by the step of the deglutathionylation reaction for which glutathionyl specificity has been demonstrated (see Scheme 1.2A, p. 111). Step 1 indicates selectivity for glutathionyl mixed disulfide substrates and/or for selective attack of Grx on the sulfur of the glutathionyl moiety. Step 2 indicates selectivity for GSH as the second substrate to reduce the Grx-SSG intermediate. ND, not determined. N/A, not applicable.

*Inferred from studies of null and multicopy mutant strains

**Studies of null mutant strains suggest a contribution to cellular deglutathionylation activity, but assays on purified, recombinant protein show little to no activity (see Section

I.D).

***Activity towards HEDS was GSH-dependent (Sun et al., 2005), but it was not distinguished whether this dependence reflected a requirement for forming a glutathionyl mixed disulfide first substrate (βME-SSG) from the pro-substrate (HEDS), for using

GSH as the preferred second substrate, or both.

119

Table 1.3 Potential phosphorylation sites of human Grx enzymes. FASTA sequences of

human Grx1, Grx2, and Grx5 were entered into NetPhos 2.0 (Blom et al., 1999) and

NetPhosK 1.0 (Blom et al., 2004) search engines to identify potential phosphorylation

sites. Sites with probability scores ≥0.6 are included. DNAPK, DNA -dependent protein kinase; PKC, protein kinase C; PKG, cyclic GMP-dependent protein kinase.

120

Grx Residue Probabilit †Location (relative isoform (FASTA) Kinase y score to active site) hGrx1 S34 DNAPK 0.6 opposite end of α-helix 2

S88 PKC 0.66 neighboring α-helix 4 hGrx2 S16 general 0.73 *

S20 general, PKA 0.91, 0.68 *

S39 general 0.69 *

S48 general 0.99 *

S73 PKC 0.61 facing active site (β-sheet 1) immediately following active T81 PKC 0.8 site (α-helix 2)

opposite side of protein Y113 general 0.73 (α-helix 3)

neighboring α-helix T137 general, PKC 0.9, 0.84 (α-helix 4)

S158 general, PKC 0.9, 0.84 *

hGrx5 S41 general 0.69 **

S156 general 0.87 **

†The hydroxyl groups of all of the indicated residues are solvent-exposed and outward- facing, with the exception of Grx2 T137, which faces the protein interior

* not included in published x-ray structure

**no structural information available

121

Table 1.4. Cardiovascular proteins regulated by S-glutathionylation.

Effect(s) of Protein Reference Function glutathionylation Reversed by GRx? ↓ polymerization rate; (Chen and myocyte actin ↓cooperativity in not shown Ogut, 2006) contraction binding tropomyosin

(Eaton et glycolysis; ↓ GAP dehydrogenase GAPDH not shown al., 2002) apoptosis activity

↓ rotenone-sensitive (Taylor et mitochondrial - Complex I activity; ↑ O2 not shown al., 2003) respiration production

(Beer et al., ↓ Complex I activity in vitro by GRx2 2004)

↑ electron transfer (Chen et mitochondrial Complex II* efficiency; not shown al., 2007) respiration ↓ electron leakage

(Adachi et cytosolic Ca2+ SERCA al., 2004b) ↑ Ca2+ uptake activity not shown reuptake **

(Sanchez et SR Ca2+ ↑ Ca2+ release rates RyR not shown al., 2005) release (transient)

↑ Ca2+ release rates (Sanchez et (transient); ↓ Ca2+ leak; al., 2008) ↓ infarct size not shown

122

Effect(s) of Protein Reference Function glutathionylation Reversed by GRx? (Adachi et al., hypertrophic ↑ protein synthesis; ↑p- via GRx1 Ras 2004a) ** signaling p38, p-Akt overexpression

↑ Ras-Raf binding; ↑ (Pimentel et Ras-GTP binding; ↑ via GRx1

al., 2006) ** ERK activation; ↑ overexpression protein synthesis

↑ Ras membrane activation of translocation; ↑ ERK- (Clavreul et MEK/ERK/Ak P; ↑ Akt-P; ↑ guanine not shown al., 2006a) t nucleotide exchange (in vitro)

↑ ERK-P (sustained); ↑ (Clavreul et insulin Akt-P (transient); ↓ via GRx1 al., 2006b) resistance insulin-induced Akt-P; overexpression ↑IRS-P

(Nulton- ↓ AKG dehydrogenase α-KGDH Persson et al., TCA cycle via GRx1 activity 2003)

(Applegate et ↓ AKG dehydrogenase via GRx1 al., 2008) activity

*Unusual example of a protein that is glutathionylated at baseline but deglutathionylated during oxidative stress (e.g., IR) **Excellent demonstrations of glutathionylation as a regulatory mechanism ***Glutathionylation achieved under supraphysiological oxidant concentrations

123

Table 1.5. Evidence for regulation of apoptosis by Grx isoforms. Summary of studies examining the potential role of Grx1 and Grx2 as regulators of apoptosis.

References are presented in chronological order. The role of each Grx isoform in apoptotic regulation, according to each study, is underlined in the “Conclusion” column.

*Limitation: lack of detail in description of apoptotic assessment makes it difficult to evaluate whether apoptosis was evaluated at a sufficient time point after MI to allow detection.

Reference Observation Proposed mechanism Conclusion (Chrestens Grx inhibition via Cd2+ ↑ glutathionylation of an Grx protects en et al., treatment  ↑ H2O2- apoptotic mediator lymphocytes from 2000) induced apoptosis in H9 H2O2-induced cells apoptosis

(Murata et Grx1 overexpression  “redox regulation” Grx1 protects al., 2003) ↓H2O2-induced of Akt cardiomyocytes apoptosis in H9c2 cells from H2O2-induced apoptosis

(Lillig et Grx2 knock-down  ↑ N/A Grx2 protects HeLa al., 2004) Dox- and PAO-induced cells from oxidant- apoptosis in HeLa cells induced apoptosis

(Enoksson Grx2 overexpression in ↓ cytochrome c release, Grx2 protects HeLa et al., HeLa cells  ↓ Dox- caspase activation, release cells from oxidant- 2005) and 2-deoxyglucose- of cardiolipin induced apoptosis induced apoptosis in HeLa cells

(Qanungo Grx1 knock-down  ↓ Grx1-mediated Grx1 sensitizes et al., apoptosis in hypoxic glutathionylation MIA PaCa cells to 2007) pancreatic cancer cells (inhibition) of apoptosis NFκB p65

(Pan and Grx1 knock-down  ↓ Grx1-mediated Grx1 sensitizes Berk, TNFα/CHX-induced deglutathionylation BAECs to 2007) apoptosis (activation) of TNFα/CHX- pro-caspase 3 induced apoptosis

124

Reference Observation Proposed mechanism Conclusion (Ho et al., MEFs from Grx1 KO Referenced Grx1 sensitizes 2007) mouse exhibit ↓ Pan and Berk, 2007 MEFs to TNFα- TNFα/actinomycin D- induced apoptosis induced apoptosis

(Malik et ND in cardiomyocyte N/A Grx1 does not al., 2008) apoptosis ± MI in Grx1- regulate apoptosis transgenic or KO mice following MI apoptosis in rodent heart*

(Nagy et Grx2-TG mice exhibit ↓ Activation of PI3K/Akt, Grx2 protects al., 2008) apoptosis following MI NFκB, and Bcl-2 cardiomyocytes in isolated working from post-MI hearts apoptosis

(Anathy et Grx1 siRNA → ↑FasL- FasL → ↑ caspase activity Grx1 protects lung al., 2009) induced apoptosis in → ↑ Grx1 degradation → epithelial cells from lung epithelial cells ↑Fas-SSG → ↑ DISC FasL-induced formation apoptosis

125

Table 1.6. Proposed functions of Bcl-2 and Bcl-xL proteins.

Function Bcl-2 Bcl-xL Reference Heterodimerization with proapoptotic Bcl-   (Chao et al., 1995; 2 family members (Bax, Bad) Gustafsson and Gottlieb, 2007)

Regulation of mitochondrial membrane   (Shimizu et al., potential 1998; Vander Heiden et al., 1999)

Inhibition of caspase activation   (Krebs et al., 1999; Bcl-2: capsase 3 Wang et al., 2004; Bcl-xL: caspases 7-9 Naumann et al., 2004; Pan et al., 1998; Hu et al., 1998)

Regulation of GSH homeostasis/redox  (Kane et al., 1993; balance Webster et al., 2006)

Regulation of calcium homeostasis via  (Rong et al., 2008)

IP3R

Inhibition of DISC formation  (Wang et al., 2004) Inhibition of PARP processing  (Naumann et al., 2004)

126

Figure 1.1. Inactivation of PTP1B by QSOX/GSH: PTP1B (2.5 μM) was incubated in

PBS in the presence of GSH (1 mM) and avian QSOX (0.05 mg/mL) at room

temperature. After various incubation times, an aliquot of the reaction mixture was

removed and assayed for phosphatase activity by dilution 1:10 into assay buffer

containing imidazole (50 mM, pH 6.8), NaCl (150 mM), and EDTA (1 mM). Assays

were initiated by addition of para-nitrophenylphosphate (pNPP, 1 mM), and rates of

pNPP dephosphorylation were determined by monitoring production of p-nitrophenol via

ΔA405 in a microwell plate reader. Closed circles () correspond to reaction mixtures

without GSH or QSOX. Closed squares () correspond to reaction mixtures containing all components. Data generated by David Starke (Gallogly and Mieyal, 2007).

127

Figure 1.2. Grx residues reported to interact with a covalently bound GS-moiety in the Grx-SSG mixed disulfide. Grx residues reported to interact with a covalently bound

GS-moiety in the Grx-SSG mixed disulfide. Residues identified as making ion pair and/or H bond contacts to the bound GS-moiety are indicated next to the chemical group with which they interact. The functionality of the interacting amino acid is indicated parentheses (guan, guanidino group). Colors indicate the species from which the Grx-

SSG structure was determined (red, E. coli Grx1 (Bushweller et al., 1994); brown, E. coli

Grx3 (Nordstrand et al., 1999; Elgan and Berndt, 2008); green, yeast Grx1 (Yu et al.,

2008; Hakansson and Winther, 2007); orange, yeast Grx2 (Discola et al., 2008), blue, human Grx1 (Yang et al., 1998)). For human Grx2 (represented in purple, (Johansson et al., 2007)), the depicted interactions correspond to those identified in a co-crystallized complex of reduced hGrx2 and GSH rather than the Grx2-SSG mixed disulfide. (*) indicates a modified orientation of the γ-glutamyl group of the associated glutathionyl moiety, in comparison to the schematic representation (adapted from Nikkola et al.

(1991)).

128

129

Figure 1.3. Predicted phosphorylation sites for human Grx isoforms. General

(NetPhos 2.0 (Blom et al., 1999)) and kinase-specific (NetPhosK 1.0 (Blom et al., 2004)) phosphorylation sites were predicted using FASTA sequences of human Grx1, Grx2, and

Grx5. All sites with probability scores >0.6 are boxed. Sites with probability scores >0.8 are marked with an asterisk (*). The CXXC active site sequence is underlined. The sequence alignment was adapted from Johansson et al. (2007).

130

131

Figure 1.4. Proposed interactions between Grx1 and cytosolic enzymes. A. Reduced

Grx1 is shown bound to ASK1 in MCF-7/ADR (adriamycin-resistant breast cancer) cells

under resting conditions (Song et al., 2002), Upon glucose deprivation, the GSH:GSSG

ratio decreases, resulting in oxidation of thiols in the Grx1 active site and disruption of its

association with ASK1. Dissociation of Grx1 and ASK1 leads to ASK1 activation and

increased apoptosis. B, Grx1 and caspase-3 are shown associated under non-stressed conditions in BAECs. TNFα-induced activation of Grx leads to pro-caspase-3

deglutathionylation, release from the complex, and cleavage by caspase-8 to active

caspase-3, triggering apoptosis (Pan and Berk, 2007).

132

Figure 1.5. Examples of acute regulation of Grx1. A, Treatment of NIH 3T3 fibroblasts with FGF resulted in robust deglutathionylation of actin within 15 minutes

(Wang et al., 2003). Actin deglutathionylation and associated cytoskeletal changes were blocked by knock-down of Grx1. B, Bovine aortic endothelial cells (BAECs) exposed to shear stress in culture exhibited increased Grx activity within the first 5-10 minutes of treatment (Wang et al., 2007). Increased Grx activity was correlated to increases in phosphorylation of eNOS and Akt. Changes in Grx activity were blocked by treatment with BCNU, while the changes in downstream events were blocked by Grx1 knock-down or transfection with catalytically inactive mutant enzyme. C, Treatment of BAECs with

TNF-α and cycloheximide was correlated with increased Grx activity within 3-6 hours, deglutathionylation of pro-caspase-3, increased caspase-3 activity, and increased apoptosis (Pan and Berk, 2007). Events downstream of caspase-3 deglutathionylation were blocked by Grx1 knock-down, or transfection of a catalytically inactive mutant.

133

134

Figure 1.6. Downstream effects of Ras glutathionylation in response to endogenous

H2O2 production. Two independent modes of Ras activation by glutathionylation are depicted here. On the left is shown how Ras-dependent and -independent pathways contribute to angiotensin II-induced hypertrophy in vascular smooth muscle cells (Adachi et al., 2004a), namely (1) coupling of angiotensin II receptor activation to production of

ROS by NADPH oxidase, followed by Ras glutathionylation and activation of Akt and p38, and (2) ROS-independent transactivation of EGFR and activation of the ERK signaling pathway. The right-hand scheme depicts a mechanism by which Ras-SSG mediates the hypertrophic response of cardiomyocytes to mechanical strain. Strain- stimulated cardiac myocytes exhibit ROS-dependent Ras glutathionylation, which activates the ERK pathway and results in increased protein synthesis. The basis for activation of Akt and p38 (left) vs. ERK (right) pathways by the same signaling intermediate (i.e., Ras-SSG) is not yet understood.

135

Figure 1.7. Downstream effects of Ras glutathionylation in response to exogenous

peroxynitrite or oxLDL. This figure depicts distinct downstream events resulting from

Ras glutathionylation in bovine aortic endothelial cells in response to peroxinitrite added exogenously (left) or generated endogenously in response to oxLDL exposure (right) as reported by Clavreul et al. (2006a and b). In response to ONOO- treatment, Ras

glutathionylation leads to transient phosphorylation of ERK and Akt; however,

subsequent downstream signaling events remain unknown. oxLDL exposure (which

leads to ONOO- production in situ) also caused Ras-SSG-dependent, transient Akt

phosphorylation (center); however, it induced a sustained time course of ERK

phosphorylation (right) as well as diminished Akt activation in response to a subsequent

stimulus (insulin). The decreased insulin-induced Akt activation conferred by oxLDL

pretreatmentcould be explained by ERK-induced phosphorylation (and inactivation) of

IRS, which is upstream of Akt in the insulin signaling pathway.

136

CHAPTER 2: HYPOTHESES, AIMS, EXPERIMENTAL APPROACHES, AND OVERVIEW OF FINDINGS

2.1. KINETIC CHARACTERIZATION OF MAMMALIAN GRX2

As described in Chapter 1, human Grx1 is the most extensively characterized glutaredoxin enzyme. Previous work in the Mieyal laboratory led to the discovery of its nucleophilic, double-displacement (ping-pong) catalytic mechanism with high commitment to catalysis (Gravina and Mieyal, 1993), as well as the basis for its rate enhancement over uncatalyzed deglutathionylation (Srinivasan et al., 1997). Many recent publications have focused on the physiological roles of Grx1, and identified potential functions in the regulation of metabolism (Eaton et al., 2002; Eaton and

Shattock, 2002), protein synthesis (Adachi et al., 2004a), calcium homeostasis (Adachi et al., 2004b), 2004), inflammation (Reynaert et al., 2006; Shelton et al., 2007; Shelton et al., 2009), and cell fate (Murata et al., 2003; Pan and Berk, 2007; Qanungo et al., 2007;

Anathy et al., 2009).

In 2001, two research groups independently reported the discovery and preliminary characterization of a second mammalian glutaredoxin, named Grx2 (Lundberg et al.,

2001; Gladyshev et al., 2001). Both groups reported that the specific activity of Grx2 towards the pro-substrate HEDS was substantially (i.e., ~10-fold) lower than that of

Grx1, and both presented evidence for mitochondrial localization of the enzyme.

Subsequent studies on human Grx2 (hGrx2) led to the surprising conclusion that unlike

Grx1, Grx2 exhibited “high affinity” for both RSSG and GSH substrates. First,

Johansson et al. (2004) interpreted the 5-6-fold lower KM,app observed for hGrx2 vs. Grx1

137

towards BSA-SSG and RNase-SSG substrates as evidence for high affinity for protein-

SSGs. Later, Lundberg et al. (2006) reported that hGrx2, unlike hGrx1, could be selectively purified using GSH-agarose beads, suggesting that it binds tightly to GSH.

High affinity of Grx2 towards either RSSG or GSH would suggest a distinction from the behavior of hGrx1, which exhibits essentially no reversible binding to RSSG or GSH

(Gravina and Mieyal, 1993). We reasoned that high-affinity interactions between Grx2 and either substrate would be reflected in the true KM values for each substrate as determined by 2-substrate kinetic analysis. Preliminary kinetic studies by David Starke suggested that Grx2, like Grx1, exhibited true KM values towards both substrates that

approach infinity, challenging the conclusion that the enzyme binds tightly to either

substrate. These controversial results, in combination with unanswered questions about

the kinetic behavior of hGrx2, guided our approach to the studies described in Chapter 3.

To identify the appropriate substrates for kinetic analysis of Grx2, we first asked

whether Grx2, like Grx1 (Gravina and Mieyal, 1993), was selective for protein-SSG

mixed disulfides. In assays containing GSH and BSA-SSG or BSA-SSC, hGrx2

enhanced deglutathionylation over the background rate only for BSA-SSG, indicating exquisite selectivity for GSH-containing mixed disulfide substrates. Next, we investigated the catalytic mechanism of Grx2, which was not yet reported in the literature. Both human and mouse Grx2 were subjected to two-substrate kinetic analysis in which CSSG and GSH were varied at fixed concentrations of the other substrate.

Then, the pattern of lines on double-reciprocal plots (1/v vs. 1/[S]) was used to distinguish between sequential and ping-pong catalytic mechanisms. These experiments indicated that Grx2, like Grx1, operates via a double displacement (ping-pong) catalytic

138 mechanism, and they also provided the data necessary to assess whether true KM values could be calculated for each substrate. (Secondary plots of 1/KM vs. 1/[S] yield true KM as the negative reciprocal of the x-intercept; projection to the origin is indicative of an essentially irreversible encounter complex). Consistent with preliminary data, true KMs for RSSG and GSH approached infinity for both human and mouse Grx2, suggesting high commitment to catalysis with no reversible binding step. The ability of Grx2 to bind

GSH was investigated further by testing the GS-analog, S-methylglutathione (GSMe) as a competitive inhibitor in the spectrophotometric assay. As shown previously for hGrx1

(Srinivasan et al., 1997), GSMe did not inhibit the Grx2-catalyzed reaction, even at millimolar concentrations.

Having verified the catalytic mechanism of Grx2, we next considered the basis for decreased specific activity of hGrx2 reported by Lundberg et al. (2001) and Gladyshev et al. (2001), and confirmed in our laboratory by Susan Ospina (see Table 3.3, p. 188). As described in Chapter 1, the rate enhancement of hGrx1 over uncatalyzed deglutathionylation can be explained by the low pKa of its catalytic Cys (the leaving group in the rate-determining step) and its special enhancement of the nucleophilicity of

GSH as the second substrate (Srinivasan et al., 1997). We hypothesized that the decrease in specific activity of hGrx2 was due to decreased pKa of its catalytic cysteine, decreased enhancement of glutathionyl nucleophilicity, or both. Before addressing these possibilities, it was important to confirm that the rate-determining step of catalysis by

Grx2 was reduction of Grx2-SSG by GSH; if not, then reasons for the decreased specific activity of Grx2 would reflect molecular events in the first step of the reaction (i.e., nucleophilic attack of Grx2 on the RSSG substrate). The pH rate profile of hGrx2-

139

catalyzed deglutathionylation indicated an inflection point around pH 8.5, reflecting the

titration of GSH and indicating that the second step of the reaction is rate-determining, as for hGrx1 (Srinivasan et al., 1997).

The pKa of hGrx2’s active site cysteine was determined by measuring the pH-

dependence of its inactivation by iodoacetamide (IAM). The inflection point of the %

inactivation vs. pH curve occurred around pH 4.5, indicating a 1 pH-unit increase in pKa

vs. Grx1 and accounting for about 40% of the 10-fold decrease in specific activity between the two isozymes. The ability of hGrx2 to enhance the nucleophilicity of GSH was addressed by determining second-order rate constants for deglutathionylation in the presence of either GSH or cysteinyl-glycine, a dipeptide subset of GSH whose nucleophilicity is not enhanced by hGrx1 (Srinivasan et al., 1997). Comparison of second-order rate constants for both reactions indicated that the enhancement of GSH nucleophilicity by hGrx2 is approximately one-half of that by hGrx1, essentially accounting for the remainder of the difference in specific activity between the two enzymes.

In addition to deglutathionylation, hGrx1 promotes glutathione thiyl radical (GS)

scavenging, as well as protein glutathionylation in the presence of activated GS-

derivatives, such as GSNO and GS (Starke et al., 2003). We reasoned that catalysis of

redox reactions using the substrate GS could represent a significant physiological

activity of hGrx2 considering its mitochondrial localization. Thus, we determined its

GS scavenging activity and measured its glutathionylating activity towards GAPDH, a

physiological target of glutathionylation (Eaton et al., 2002) in the presence of GSSG,

GSNO, and GS. GS-scavenging activity by hGrx2 was ~10-fold slower than that of

140

hGrx1, suggesting that the final proposed step of catalysis (deglutathionylation of the

Grx-SSG intermediate (Starke et al., 2003) is rate-determining, identical to the rate-

determining step in deglutathionylation). Glutathionylation with GSSG was ~4-fold

slower for hGrx2 vs. hGrx1, consistent with a catalytic mechanism that is the reverse of

deglutathionylation, with reduction of Grx-SSG by protein-SH being the rate-determining step (and rate enhancement determined only by the difference in the pKa of the catalytic

Cys). GAPDH glutathionylation by GS was enhanced equally by hGrx1 and hGrx2,

supporting a mechanism in which catalytic cysteine pKa and glutathionyl nucleophilicity

do not influence the speed of the rate-determining step in this reaction.

Concurrent with our kinetic studies, Dr. Harish Pai demonstrated that Grx1 and Grx2

exhibit distinct submitochondrial localizations in rat heart and liver (Pai et al., 2007), and

he also determined the relative quantities of each enzyme (i.e., ng per mg mitochondrial

protein) in each mitochondrial subcompartment. These determinations allowed us to

place our kinetic observations about mammalian Grx2 into a physiological context by

predicting the relative contributions of each Grx isoform to total mitochondrial

deglutathionylating (and glutathionylating) activity, using specific activity measurements,

Grx concentrations, and estimated protein concentration and pH of each mitochondrial

compartment (see Chapter 3). Our prediction of comparable deglutathionylation

activities in the mitochondrial intermembrane space must be verified by analysis of total

deglutathionylation activity in extracts from each mitochondrial subcompartment (see

Chapter 5).

141

2.2. REGULATION OF APOPTOSIS IN CARDIOMYOCYTES BY GRX1 VIA

NFΚB AND TARGET GENES BCL-2 AND BCL-XL

Three sets of observations led us to investigate the role of Grx1 as a regulator of apoptosis in the aging heart. First, increased susceptibility to apoptosis appears to be a contributing factor to the increased cardiac injury experienced in elderly human patients and animals following myocardial infarction (MI). This premise arises from observations that aging is an independent risk factor for post-MI injury (Rich et al., 1992; Lesnefsky et al., 1996), apoptosis contributes significantly to post-MI injury (Mani, 2008; Zidar et al.,

2007; Lee and Gustafsson, 2009), and cardiomyocytes from elderly animals exhibit increased apoptosis both at baseline and following an oxidative challenge (Frolkis et al.,

1991; Tani et al., 1997; Ashton et al., 2006; Willems et al., 2005).

Second, recent work highlights glutaredoxin as a regulator of cell fate decisions in cardiomyocytes (Murata et al., 2003), fibroblasts (Ho et al., 2007), T cells (Chrestensen et al., 2000), pancreatic cancer cells (Qanungo et al., 2007), lung epithelial cells (Anathy et al., 2009), and endothelial cells (Pan and Berk, 2007). Moreover, some groups have demonstrated specifically that Grx-mediated deglutathionylation of apoptotic mediators affects their activities (e.g., pro-caspase 3 (Pan and Berk, 2007), p65 (Qanungo et al.,

2007), IKK (Reynaert et al., 2006)).

Third, previous work by David Starke, Dr. Carol Chrestensen, and Tyler Murphy

(Mieyal Laboratory) indicated that Grx1 content and total Grx activity were decreased by approximately 40% with aging in cytosolic fractions of heart tissue isolated from Fischer

344 rats, while other TDOR enzymes remained unchanged (see Chapter 4). Based on the important contributory role of apoptosis to cardiac injury, the ability of Grx1 to regulate

142

apoptosis in cardiomyocytes, and the selective diminution in Grx1 in the aging heart, we

hypothesized that the age-related decreases in Grx1 contribute to the increased apoptotic

susceptibility of cardiomyocytes from elderly animals. Therefore, subsequent studies

focused on the question of whether diminished levels of Grx1 increase apoptotic

susceptibility in cardiomyocytes, and if so, by what mechanism.

To determine whether diminution of Grx1 was sufficient to increase apoptotic

susceptibility in cardiomyocytes, we knocked down Grx1 in H9c2 cells (rat embryonic

cardiomyocytes) to a similar extent as observed with aging (i.e., ~50%) and measured

apoptosis in response to H2O2 treatment (Han et al., 2004) and simulated IR (Kim et al.,

2006). Two methods of Grx1 knock-down were employed: transient knock-down using siRNA transfection and stable knock-down using an adenovirus containing shRNA directed towards Grx (see Chapter 4, Materials and Methods). For both methods of knock-down, Grx1 deficiency led to increased apoptosis at baseline and following an oxidative challenge. This work represents the first time that the effect of diminished

Grx1 on apoptosis was evaluated in this cell line, and our observations complement and extend the report of Murata et al. (2003), who found that overexpression of Grx1 in H9c2 cells led to resistance to oxidant-induced apoptosis.

Grx1 has been implicated in the regulation of apoptosis via several mechanisms, including redox regulation of Akt (Hirota et al., 2000; Murata et al., 2003), reversible glutathionylation of NFκB-p65 (Qanungo et al., 2007), Fas (Anathy et al., 2009), and/or procaspase-3 (Pan and Berk, 2007), and protein-protein interaction with ASK1 (Song et al., 2002). Another NFκB pathway component, IKK, was recently shown to be regulated by reversible glutathionylation and by Grx1 (Reynaert et al., 2006), although the effect of

143

this regulation on apoptosis has not yet been determined. We hypothesized that the most likely mechanism of apoptotic regulation by Grx1 in H9c2 cells was via regulation of the

NFκB pathway, most likely through deglutathionylation, for several reasons. First, multiple members of the NFκB pathway are regulated by reversible glutathionylation

(Figure 2.1, p. 149, reviewed in Shelton and Mieyal (2008), with glutathionylation being inhibitory in nearly every case. Thus, diminution of Grx1 would be expected to lead to increased glutathionylation of the affected pathway members, decreasing total transcriptional activity. Second, pilot experiments conducted by Dr. Harish Pai demonstrated that (1) H9c2 cells with stable Grx1 knock-down exhibited decreased basal

NFκB activity, and (2) inhibition of NFκB activity in wild-type H9c2 cells sensitized

them to apoptosis. Finally, regulation of apoptosis by deglutathionylation (activation) of

pro-apoptotic molecules (i.e., procaspase-3) would be inconsistent with the observation

that Grx1 knockdown increases apoptotic susceptibility.

Dr. Melissa Shelton (Mieyal laboratory) measured NFκB activity in control and Grx1

knock-down cells using a Luciferase reporter assay. The measurement of NFκB activity

was coordinated with the time course of Grx1 knock-down so that transcriptional activity

was measured precisely at the time in which Grx1 was knocked down to a similar extent

as observed with aging (i.e., ~50%). NFκB activity in Grx1-deficient cells was indeed diminished (by ~30%), while content of the transcription factor components p50 and p65 was not significantly different from the amount detected in control cells.

To determine whether the decrease in NFκB activity observed with Grx1 knock-down

was sufficient to increase apoptotic susceptibility in H9c2 cells, wild-type H9c2 cells were treated with BMS 345541 (Burke et al., 2003), a selective inhibitor of IKK, and

144

apoptotic susceptibility was measured in the presence and absence of H2O2. Notably, the

concentration of BMS which inhibited NFκB activity to a comparable extent as observed in Grx1 knock-down cells was also sufficient to increase apoptotic susceptibility, suggesting that the diminution of NFκB in Grx1 deficient cells contributes to their decreased threshold for apoptosis.

We hypothesized that decreased NFκB activity led to increased apoptotic

susceptibility in Grx1-deficient cells via decreased transcription of anti-apoptotic target

genes. We first focused on Bcl-2 and Bcl-xL because of their demonstrated anti-

apoptotic roles in the heart, especially in the context of oxidative insult (see Chapter 4).

mRNA and protein levels of Bcl-2 and Bcl-xL were determined by real-time PCR and

Western blotting, respectively, and were found to decrease by approximately one-half

with Grx1 knock-down. Next, the specific contributions of each protein to apoptotic susceptibility in H9c2 cells were examined by targeted knock-down studies in which one or both proteins were selectively diminished to a similar extent as observed in Grx1- deficient cells, and apoptotic susceptibility was determined with and without H2O2

exposure. Diminution of Bcl-2 or Bcl-xL increased apoptotic susceptibility at baseline,

while knock-down of Bcl-xL but not Bcl-2 increased apoptosis following oxidative insult,

suggesting distinct anti-apoptotic roles for the two proteins in the context of oxidative

cardiac injury.

To determine whether the mechanism of apoptotic regulation by Grx1 delineated in

H9c2 cells might occur also in aging cardiomyocytes, we compared NFκB activity in

cardiomyocytes from adult (6-10-mo) and elderly (24-27-mo) Fischer 344 rats. In parallel, we confirmed that Grx activity was diminished (by ~40%) in cardiomyocytes

145

isolated from elderly rats compared to adult rats (experiments by David Starke, Mieyal

Laboratory), and verified that cardiomyocytes isolated from elderly animals exhibited the

same overall structural features (e.g., rod shape, characteristic striations) as those from

adults. As observed in Grx1 knock-down H9c2 cells, cardiomyocytes from elderly rats

(i.e., Grx1-deficient primary cardiomycoytes) exhibited decreased NFκB activity

compared to those from adult rats, consistent with the proposed mechanism of Grx1-

mediated regulation of apoptosis demonstrated in H9c2 cells (Scheme 2.1, p. 148).

Further confirmatory studies in primary cardiomyocytes are proposed in Chapter 5.

2.2.1 JUSTIFICATION OF THE MODEL SYSTEMS

Two model systems were utilized for studies of apoptotic regulation by Grx1 in

cardiomyocytes: H9c2 cells and adult (~6-mo) and elderly (2~4-mo) Fischer 344 (F344)

rats. F344 rats are an inbred, albino rat strain maintained in homogenous, well-controlled populations by the National Institute of Aging. Elderly F344 rats exhibit hallmarks of human cardiovascular aging, including exacerbated post-MI injury (Lesnefsky et al.,

1994); and they represent a commonly used animal model for the investigation of age- related changes in the cardiovascular system (Caffrey et al., 1994; Isoyama et al., 1987;

Phaneuf and Leeuwenburgh, 2002). Previous work by David Starke (Mieyal laboratory) indicated that Grx1 content and activity were diminished with aging in cytosolic fractions of heart tissue from elderly vs. adult F344 rats. We hypothesized that the age-related decrease in Grx1 with aging contributes to the increased apoptotic susceptibility of cardiomyocytes from elderly animals (Kajstura et al., 1996; Nitahara et al., 1998; Liu et al., 2002). Aging is a complex phenotype involving many molecular changes (see

Juhaszova et al., 2005), so to investigate the specific role of diminished Grx1 on

146

apoptotic susceptibility, we conducted mechanistic studies in H9c2 cells (summarized

above and presented in detail below), and then used primary cardiomyocytes from adult

and elderly F344 rats for confirmatory experiments (e.g., NFκB activity, see Chapter 4).

H9c2 cells are an immortalized rat embryonic cardiomyocyte cell line used routinely

to study redox signaling mechanisms in cardiomyocytes (Murata et al., 2003; Urata et al.,

2006; Kanda et al., 2006). H9c2 cells grow relatively quickly (doubling time ~2 days) and are easily manipulated to achieve targeted overexpression or knock-down of a protein of interest. In the studies described in Chapter 4, H9c2 cells were used to investigate the specific effects of Grx1 diminution on apoptotic susceptibility, NFκB activity and mRNA and protein content of the anti-apoptotic NFκB targets Bcl-2 and Bcl-xL. The effects of diminution of Bcl-2 and Bcl-xL on apoptotic susceptibility were also investigated in

H9c2 cells. Following these mechanistic studies, the effect of diminished Grx1 on NFκB activity was confirmed in cardiomyocytes isolated from adult (6-mo) and elderly (24-mo)

F344 rats.

147

Scheme 2.1. Model of Grx1-mediated regulation of apoptosis in cardiomyocytes.

We hypothesized that diminished content and activity of Grx1 with aging would lead to

increased glutathionylation of component(s) of the NFκB pathway (designated as

[NFκB]). As demonstrated in other model systems, we expected glutathionylation of one

or more NFκB pathway components (i.e., [NFκB]-SSG) to inhibit NFκB-dependent

transcription, resulting in decreased content of anti-apoptotic NFκB target genes (e.g.,

Bcl-2 and Bcl-xL). Decreased levels of Bcl-2 and Bcl-xL were expected to sensitize cardiomyocytes to apoptosis, and indeed, selective diminution of Bcl-2 or Bcl-xL

increased apoptosis at baseline in H9c2 cells. In contrast, diminution of Bcl-xL but not

Bcl-2 increased susceptibility to apoptosis triggered by H2O2 exposure (see Chapter 4).

148

Figure 2.1. Sites of regulation of the NFκB pathway by reversible glutathionylation.

Shown is the canonical NFκB pathway in which IKK (IκB kinase) phosphorylates IκB

(inhibitor of κB), leading to its ubiquitination and degradation. Degradation of IκB allow

release of the p50-p65 heterodimer, which translocates to the nucleus, binds to specific

promoter sequences, and activates transcription of anti-apoptotic target genes, enhancing survival. Sites of regulation of the NFκB pathway by reversible glutathionylation are

outlined in red and include IKK (Reynaert et al., 2006; Shelton et al., 2009), p50 (Pineda-

Molina et al., 2001), p65 (Qanungo et al., 2007), E1 and E2 ubiquitin ligases (Jahngen-

Hodge et al., 1997; Obin et al., 1998), and the 20S proteasome (Demasi et al., 2003).

NFκB pathway activator Ras (Adachi et al., 2004a) and inhibitor PTEN (Cruz et al.,

2007) are also regulated by reversible glutathionylation. The redox status and activity of

Akt, which has been shown to activate the NFκB pathway, is regulated by Grx1 (Murata et al., 2003), but the specific mechanism of redox regulation (e.g., reversible glutathionylation) is unresolved.

149

CHAPTER 3: KINETIC AND MECHANISTIC CHARACTERIZATION AND VERSATILE CATALYTIC PROPERTIES OF MAMMALIAN GLUTAREDOXIN 2: IMPLICATIONS FOR INTRACELLULAR ROLES

The contents of this chapter represent an expansion from an article of the same title published in Biochemistry (47(42):11144-57). Reproduced with permission from the American Chemical Society (Copyright 2008).

3.1 Abstract

Glutaredoxin (Grx)-catalyzed deglutathionylation of protein-glutathione mixed disulfides

(protein-SSG) serves important roles in redox homeostasis and signal transduction,

regulating diverse physiological and pathophysiological events. Mammalian cells have

two Grx isoforms: Grx1, localized to the cytosol and mitochondrial intermembrane

space, and Grx2, localized primarily to the mitochondrial matrix (Pai et al., 2007). The

catalytic behavior of Grx1 has been characterized extensively; whereas Grx2 catalysis is

less well understood. We observed that human Grx1 and Grx2 exhibit key catalytic

similarities, including selectivity for protein-SSG substrates and a nucleophilic, double- displacement, monothiol mechanism exhibiting high commitment to catalysis. A key distinction between Grx1- and Grx2-mediated deglutathionylation is decreased catalytic efficiency (kcat/KM) of Grx2 for protein deglutathionylation (due primarily to decreased

kcat,), reflecting a higher pKa of its catalytic cysteine, as well as a decreased enhancement

of nucleophilicity of the second substrate, GSH. As documented previously for hGrx1

(Starke et al., 2003), hGrx2 catalyzes glutathione-thiyl radical (GS•) scavenging; and it

also mediates GS-transfer (protein S-glutathionylation) reactions, where GS• serves as a superior glutathionyl-donor substrate for formation of GAPDH-SSG, compared to GSNO and GSSG. In contrast to its lower kcat for deglutathionylation reactions, Grx2 promotes

150

GS-transfer to the model protein substrate GAPDH at rates equivalent to those of Grx1.

Estimation of Grx1 and Grx2 concentrations within mitochondria predicts comparable deglutathionylation activities within the mitochondrial subcompartments, suggesting localized regulatory functions for both isozymes.

3.2 Introduction

Protein glutathionylation, or mixed disulfide formation between a protein cysteine moiety and the cysteine moiety of glutathione (GSH), is a post-translational modification with important roles in cellular signal transduction and sulfhydryl homeostasis within mammalian cells (Klatt and Lamas, 2000; Shelton et al., 2005; Gallogly and Mieyal,

2007). During overt oxidative stress, glutathionylation is thought to protect critical sulfhydryl groups from irreversible oxidation to sulfinic and sulfonic acids (Shelton et al.,

2005; Thomas et al., 1995; Mieyal et al., 1995). Under non-stressed conditions, reversible glutathionylation is a redox signal transduction mechanism that modulates protein activity, either through activation or inactivation, thereby regulating processes such as protein synthesis (Adachi et al., 2004a; Pimentel et al., 2006), calcium homeostasis (Adachi et al., 2004b), cell growth (Barrett et al., 1999b), and transcription factor activity (Qanungo et al., 2007; Reynaert et al., 2006). Although mechanisms for protein glutathionylation are not fully understood (Gallogly and Mieyal, 2007), multiple studies indicate that within mammalian cells, protein deglutathionylation is catalyzed principally by the thiol-disulfide oxidoreductase enzyme glutaredoxin (Grx, aka thioltransferase) (Chrestensen et al., 2000; Jung and Thomas, 1996; Ho et al., 2007).

Mammalian glutaredoxin (Grx1), localized to the cytosol and mitochondrial intermembrane space (Pai et al., 2007), is well characterized as a specific and efficient

151

catalyst of deglutathionylation of protein-SSG, proceeding via a nucleophilic, double-

displacement mechanism in which the N-terminal Cys (Cys22) of the CPY(F)C active

site attacks the glutathionyl sulfur of the protein-SSG disulfide bond, releasing protein-

SH and forming a Grx1-SSG covalent intermediate (Scheme 3.1A, p. 184; Gravina and

Mieyal, 1993). In the second, rate-determining step of the reaction, GSH serves as the

nucleophile in the thiol-disulfide exchange with Grx1-SSG, forming Grx1-S- and GSSG

(Gravina and Mieyal, 1993; Srinivasan et al., 1997). Although it contains vicinal thiols

in its active site (a common property of the thiol-disulfide oxidoreductase (TDOR) enzymes) hGrx1 is distinguished from other TDORs, such as thioredoxin (Trx), in this monothiol mechanism, i.e., only the N-terminal active site Cys is required for catalysis

(Yang et al., 1998). In fact, the presence of the C-terminal active site Cys limits kcat by

promoting a side reaction whereby the Grx1-SSG intermediate is converted to an

intramolecular disulfide (Scheme 3.1A, p. 184) that can be recruited back into the

catalytic cycle by reaction with GSH. Rate enhancement of the Grx1-catalyzed reaction

over the non-enzymatic reaction is attributed primarily to the superior ability of the active site thiolate to act as a leaving group in the second step of the reaction (Srinivasan et al.,

1997), due to its unusually low pKa of 3.5 (Mieyal et al., 1991b; Gan and Wells, 1987).

In addition, Grx1 appears to enhance the nucleophilicity of GSH for the second step of

the reaction (Srinivasan et al., 1997) (Scheme 3.1A, p. 184).

A second mammalian glutaredoxin isoform (Grx2) was discovered more recently by

homology search analysis of human and rodent EST libraries (Gladyshev et al., 2001;

Lundberg et al., 2001). Although it exhibits <35% sequence homology with Grx1, Grx2 shares distinguishing structural features, including a thioredoxin fold, a CXXC active site

152 motif (CSYC in Grx2 vs. CPYC in Grx1), and conserved amino acids implicated in the stabilization of the adducted glutathionyl moiety (Yang et al., 1998; Lundberg et al.,

2001; Gladyshev et al., 2001; Sun et al., 1998). To date, three sub-forms of Grx2

(Grx2a,b,c) have been described that appear to result from alternative splicing of the gene’s first exon (Gladyshev et al., 2001; Lundberg et al., 2001; Lonn et al., 2007), resulting in distinct N-terminal sequences. The N-terminus of Grx2a contains a mitochondrial localization sequence sufficient to target to the mitochondria either overexpressed Grx2 or a GFP fusion protein which contains the localization sequence

(Gladyshev et al., 2001). Moreover, studies of isolated rat mitochondria have documented that Grx2 is localized exclusively in the matrix, whereas Grx1 is found only in the intermembrane space of mitochondria (Pai et al., 2007). Grx2b and Grx2c share overlapping N-terminal regions transcribed from an alternatively spliced region of Exon

1 (Lonn et al., 2007). The expression pattern of Grx2b and Grx2c appears to be more restricted than that of Grx2a, being detected primarily in testes, immortalized cell lines, and tumors (Lonn et al., 2007). In contrast to Grx2a, the subcellular localization of

Grx2b and Grx2c remains unresolved, with reports of nuclear, perinuclear, and diffuse cytosolic and nuclear localizations (Lundberg et al., 2001; Lonn et al., 2007). Within mitochondria, it has been reported that Grx2a exists in an inactive dimer bridged by a

2Fe2S cluster (Lillig et al., 2005), and Grx2c is also reported to form the 2Fe2S cluster in a reconstitution assay in vitro (Lonn et al., 2007). The dimeric Grx2 complex is reported to dissociate in the presence of certain oxidants and reductants in vitro (Lillig et al.,

2005), and under conditions that promote S-ntrosylation (Hashemy et al., 2007); however, mechanisms of regulation of reversible cluster formation in vivo are unknown.

153

To date, several studies of Grx2 have emphasized apparent differences in catalytic

activity in comparison to Grx1 (Lundberg et al., 2001; Johansson et al., 2004; Hashemy et al., 2007; Fernando et al., 2006). In contrast, this report describes identical behavior for Grx2 in many of the characteristic features of the catalytic mechanism delineated previously for Grx1-mediated deglutathionylation. In particular, Grx2 displays specificity for glutathione-containing mixed disulfides, and a double displacement kinetic mechanism with high commitment to catalysis that precludes reversible binding of substrates. In addition, we observed that Grx2, like Grx1, exhibits glutathionyl-thiyl

radical (GS•)-scavenging and GS•-mediated protein S-glutathionylation activities, suggesting specific roles in redox and sulfhydryl homeostasis within the mitochondrial matrix under conditions of oxidative stress.

3.3 Materials and Methods

3.3.1 Materials

Cysteinyl-glutathione mixed disulfide was purchased from Toronto Research Chemicals,

and NADPH was from Roche. Plasmid DNA (pET-24d, Novagen) encoding human

Grx1 was generated as described by Chrestensen et al. (1995). Plasmids encoding

mature (i.e., exons 2-4, amino acids 41-164) human or mouse Grx2a were kindly

provided by Dr. Vadim Gladyshev (University of Nebraska). All other routine

chemicals were reagent grade or better, and obtained from Sigma.

Enzymes – Recombinant human Grx1 (hGrx1) and Grx2 (hGrx2) and mouse Grx2

(mGrx2) were purified as described previously (Jao et al., 2006) from their respective plasmid DNA constructs (see above) with minor modifications. Plasmid DNA (pET-

21d(+), Novagen) encoding human or mouse Grx2 was transformed into E. coli BL21

154

(DE3) competent cells. Aliquots of transformed E. coli were spread on LB plates

containing ampicillin (50 μg/mL) and incubated overnight at ºC. 37 Bacteria from a

single colony was used to inoculate 50mL LB (50 μg/mL ampicillin), which was shaken

overnight at 37ºC. A 20mL aliquot of the overnight culture was added to 1L LB broth

(50 μg/mL ampicillin), and the culture was shaken until its absorbance at 600 nm reached

1.0. Grx2 expression was then induced by adding IPTG (final concentration 1mM) to the culture. After an additional incubation for 5-6 h at 37°C, the cells were pelleted at 6000 g for 15 min at 4 °C, and the pellet was resuspended in ~15 mL 10 mM potassium phosphate buffer (pH 7.4) containing 1 mM DTT. Bacteria were lysed via French press, and the lysate was centrifuged at 10,000 g for 30 min at 4 °C. Nucleic acid concentration of the supernatant was determined by measuring A260, and 20 μg protamine sulfate was

added per 260 nm absorbance unit measured. Precipitated nucleic acids were removed by

centrifugation at 10,000 g for 30 min at 4 °C. To denature heat-sensitive proteins, 2 mL

aliquots of the supernatant were stirred in a 60 °C water bath for 1 minute, then for an

additional 30 seconds on ice. Aliquots were then pooled and precipitated proteins were

removed by centrifugation at 10,000 g for 30 min at 4 °C. The supernatant was passed over a Sephadex G-75 size exclusion chromatography column preequilibrated at 4 °C with running buffer (10 mM ammonium formate, pH 5.8). Fractions were assayed for

Grx activity using the standard spectrophotometric assay (see below), and for protein

content (BCA Assay, Pierce). Fractions with high Grx activity and low protein content

were passed over a Q-Sepharose Fast Flow anion exchange column (Pharmacia) to

remove residual nucleic acids (running buffer: 10 mM ammonium formate, pH 5.8), and

eluent was concentrated in an Amicon stirring concentrator under N2 using a 10 kDa

155

molecular weight cutoff filter at 4ºC. Glycerol (20% by volume) was added to the concentrated product and aliquots were stored at -80ºC.

Purified hGrx2 C40S protein was kindly provided by Drs. Catrine Johansson and Arne

Holmgren (Karolinska Institute, Sweden). Glutathione disulfide reductase from baker’s yeast (~170 U/mg), thioredoxin reductase from rat liver (~170 U/mg), and glutathione peroxidase from bovine erythrocytes (680 U/mg) were purchased from Sigma.

Horseradish peroxidase (~225 U/mg) was obtained from Roche.

Synthesis of [3H] BSA-SS-glutathione substrate – Synthesis was performed as described

by Chrestensen et al. (2000) with minor modifications. In brief, a 1.4-fold molar excess

of N-Succinimidyl 3-(2-pyridyldithio)-propionate (SPDP) (dissolved in water-free

(molecular sieves) dimethyl formamide) was added to 3mM S-carboxymethyl-BSA

slowly with constant stirring at room temperature in phosphate buffered saline, (PBS, 137

mM NaCl, 2.7 mM KCl, 4.3 mM sodium phosphate, 1.4 mM potassium phosphate, pH

7.4), over 1 h. The BSA derivative was then treated with 5 mM [3H] GSH (~1nCi/nmol)

for 1 h at room temperature. Excess [3H] GSH and SPDP were separated from the [3H]

BSA-SSG by Sephadex G-25 column chromatography (preequilibrated with PBS). The

protein fractions (with radiolabel) were pooled and concentrated to ~1 mM with respect

to [3H] GS-equivalents. [35S] BSA-SSG was prepared by a completely analogous

procedure, except that [35S] GSH was substituted for [3H] GSH. Both radiolabeled

substrates reacted indistinguishably in deglutathionylation assays.

Synthesis of BSA-SS-[14C] cysteine substrate – BSA-SS-[14C]cysteine (BSA-SSC) was prepared as described by Yang et al. (1998).

156

3.3.2 General Methods

Deionized, distilled water used in all enzyme assays was pre-treated with 5 g/L Chelex

resin to chelate metal cations.

3.3.3 Specific Methods

Gel electrophoresis and Western blotting – Aliquots of E. coli lysate at various stages of

Grx purification, or 2.5-80 ng isolated, purified Grx, were mixed with10 μL 4X sample

buffer (100 mL 0.5M Tris-HCl, pH 6.8, 80 mL glycerol, 160 mL 10% SDS, 20 mL 1%

bromophenol blue; final concentrations at 1X: 34 mM Tris-HCl, pH 6.8, 5.5% glycerol,

1.1% SDS, 0.014% bromophenol blue), DTT (10 mM, final) and enough PBS to reach a total volume of 40 μL. Each sample was heated to 95 °C for 10 min, then IAM was added (25 mM, final), and samples were heated for 5 min more. Samples were then loaded onto 12.5% polycacrylamide gels and resolved by electrophoresis (150 V x 1 h).

Gels were either stained using Coomassie Brilliant Blue stain (Bio-Rad) for 1 h or transferred to PVDF membranes. Membranes were blocked with 5% nonfat milk dissolved in Tris-buffered saline (TBS) for 1 h. For detection of hGrx1, blocked membranes were incubated with anti-Grx1 antibody (Steven A. Gravina, PhD thesis,

Case Western Reserve University, 1993) in 5% milk for 1 h. Blots were then washed 3 times for 10 min each with TBS and incubated with anti-rabbit secondary antibody

(Jackson Laboratories, 1:10,000 dilution), washed again, and developed using Western

Lightning chemiluminescent reagent (Perkin Elmer). For Grx2, blocked blots were incubated with anti-Grx2 antibody (gift from Dr. Vadim Gladyshev, University of

Nebraska, 5 μg/mL final) in 5% milk for 1 h, washed, incubated with fluorescein-linked, anti-rabbit secondary antibody (1:1000, Amersham) in 5% milk for 1 h, washed,

157

incubated with anti-fluorescein antibody (1:2500, Amersham) in 5% milk for 1 h, washed

again, and developed using ECF reagent (Amersham) on a Fluorimager (Bio-Rad).

Spectrophotometric Assay of Grx Activity– The coupled spectrophotometric assay for Grx activity was performed as described previously (Jao et al., 2006; Yang et al., 1998).

Reaction mixtures containing Na/K-phosphate buffer (0.1mM, pH 7.5), NADPH

(0.2mM), GSSG reductase (2 U/mL), GSH (0.02-3 mM), and Grx (0-135 nM) were prepared in microwells of a 96-well plate (final volume 200 μL) and incubated for 5 min. at 30 °C. For CSSG and BSA-SSG, reaction rates were determined within a range of concentrations from 0.25-4 times the apparent KM, while for GSH, the high non-

enzymatic rate observed at high substrate concentrations prevented determinations at

[GSH] > ~2 × KM. For the cysteine-SS-glutathione substrate (CSSG), reactions were initiated by addition of CSSG (0.002-0.5 mM); for the BSA-SS-glutathione substrate

(BSA-SSG), initial mixtures contained BSA-SSG (5-40 μM) and deglutathionylation reactions were initiated by addition of GSH (0.1-1 mM). NADPH oxidation (equivalent to GSSG formation) was monitored according to decreasing absorbance at 340 nm over 5 min (during which reaction rates remained linear with time) using a ThermoMax microplate reader (Molecular Devices). Non-enzymatic rates were subtracted from enzymatic rates and nmol NADPH oxidized/min/nmol enzyme (i.e., turnover, min-1) was

calculated using the standard extinction coefficient of NADPH (ε = 6.2 mM-1 cm-1), along with factors correcting for path length and actual NADPH absorptivity in the plate reader.

Specific activity determinations were performed in the presence of 0.1 mM CSSG and

0.5 mM GSH (units of activity per mL) was divided by the Grx protein concentration to yield U/mg. For the BSA-SS-glutathione substrate (BSA-SSG), experiments were

158

conducted as described except that BSA-SSG was substituted for CSSG. All velocity vs.

substrate plots were fit to the Michaelis-Menten equation, and apparent kinetic constants

for CSSG, BSA-SSG, and GSH were calculated accordingly. Standard error values for

kinetic constants were calculated using the Prism program (GraphPad Software).

Aliquots of standard hGrx1 protein were assayed under standard conditions periodically

for monitoring the consistency of assay solutions, temperature, and plate reader

conditions during accumulation of other kinetic data. When small deviations from the

established values for standard Grx1 activity were noted, the companion data were

normalized accordingly.

pH-dependence of Grx2 inactivation by iodoacetamide (IAM) – The pH-dependence of

inactivation of Grx2 by IAM was carried out as described previously (Jao et al., 2006;

Mieyal et al., 1991b) with minor modifications. Grx2 (3 μM) was preincubated in the

following buffers with or without IAM (0.3 mM) for 3 minutes: Na citrate, pH 3.5; Na

acetate, pH 3.5, 4.0, 4.5, 5.0; MES, pH 5.0, 5.5, 6.0. All buffers were 10 mM in

concentration, and ionic strength was adjusted to 0.5 M in all cases by addition of the

appropriate amount of NaCl or KCl. Following preincubation, Grx activity was

determined by adding an aliquot (4 μL) of the preincubation mixture to the

spectrophotometric assay system described above. Percent activity remaining was

calculated by dividing rates of deglutathionylation after preincubation with IAM by rates

in the absence of IAM and multiplying by 100%. Percent activity was plotted vs. pH and

an adaptation of the Henderson-Hasselbalch equation was used to fit the data and solve for pKa:

 10( pH − pKa)  = − ×   % activity remaining 100 100  ( pH − pKa)  1+10 

159

Relative rates of dethiolation of radiolabeled BSA mixed disulfide substrates by the Grx

isoforms – The standard radiolabel assay for Grx activity was performed as described

previously (Shelton et al., 2007; Srinivasan et al., 1997). Reaction mixtures containing

Na/K phosphate buffer (0.1 M, pH 7.5), GSH (0.5 mM), and Grx (3-480 nM) were prepared in Eppendorf tubes and placed in a 30°C water bath for 5 min. Following preincubation, reactions were initiated by addition of [3H] BSA-SSG or [14C] BSA-SS-

Cys (0.1mM final). Aliquots of the reaction mixture were removed at 0.5, 1, 2, and 3

min., and quenched by addition to an equal volume of ice cold 20% TCA. Precipitated

protein was sedimented by centrifugation at 13,000xg for 5 minutes at 4 °C, and

supernatants were analyzed for non-protein associated radioactivity (i.e. radiolabeled

GSSG or GSS-Cys). Background radioactivity contributed by non-precipitated BSA-

SSR substrate (< 0.2 % total) was measured independently and subtracted from each time

point. Enzyme-mediated dethiolation rates were determined by calculating the difference

in time-dependent radiolabel release in the presence and absence of Grx. To measure the

pH-dependence of hGrx2-catalyzed deglutathionylation (pH-rate profile), assays were

performed as described above, except that the concentration of GSH was reduced to 0.25

mM to minimize non-enzymatic rates of deglutathionylation. Also, Na/K phosphate

buffer was replaced with MES (pH 5.5-6.5), HEPES (pH 7-8), HEPPSO (pH 8.5),

glycine (pH 9-10.5). All buffers were 0.1 M, and ionic strength was adjusted to 0.3 M

using NaCl. Rates of hGrx2-dependent deglutathionylation (expressed as turnover, min-

1) were expressed as % maximal activity, and plots of activity vs. pH were fit to a

derivation of the Henderson-Hasselbalch equation:

 10( pH − pKa)  % maximal activity = ×   100  ( pH − pKa)  1+10 

160

For comparison of deglutathionylation rate constants in the presence of alternative RSH

substrates, i.e., GSH or cysteinylglycine, the concentration of RSH was 0.25 mM, and

reactions were carried out in AMPSO buffer (0.1 M, pH 9.5). Second-order rate

constants for formation of the respective R-SSG products (GSSG or Gly-CSSG) were

calculated by dividing the concentration of R-SSG produced per min by the concentration of BSA-SSG substrate for the uncatalyzed rate; or by dividing by the concentration of

Grx for the catalyzed reaction, as described previously (Srinivasan et al., 1997). In both

cases, the quotients were then divided by the concentration of RSH, and the second-order

-1 .-1 rate constants (knon-enz and kGrx) are expressed as M min .

Comparison of Grx-mediated rates of deglutathionylation of CSSG in the presence of TR

or GSH and GR – hGrx activity was determined by the spectrophotometric assay

described above with minor changes in component concentrations. All assays contained

Na/K-phosphate buffer (0.1 M, pH 7.5) and NADPH (0.2 mM), and were initiated by

addition of CSSG (0.5 mM, final). To measure catalytic rates in the presence of TR,

[hGrx1or 2] = 3.7 μM, and [TR] = 230 nM. TR activity was confirmed independently by

measuring NADPH-dependent reduction of DTNB in Na/K phosphate buffer (0.1M, pH

7), and this activity matched that the value reported by the supplier (reported = 29.4,

observed = 29.3 ± 1.9 U/mL, 1 U = Δ1 AU412/min). For measurement of turnover under

low GSH conditions, [hGrx1] was adjusted to 0.37 μM (to achieve linear dependence on

enzyme concentration), [Grx2] = 3.7 μM, [GSH] = 0.1 mM, and [GR] = 0.01 μM.

Concentrations of GSH and GR were deliberately chosen to represent one-tenth of

estimated intracellular concentrations (as described in Starke et al., 1997).

161

Glutathionyl-thiyl radical (GS•) scavenging activity of hGrx2 – GS•-scavenging activity was measured as described by Starke et al. (2003). Reaction mixtures were prepared as described for the standard spectrophotometric assay (0.1M Na/K-phosphate buffer, pH

7.5, 0.2 mM NADPH, 0.5 mM GSH, 2 U/mL GSSG reductase, and 0.01-1 μM hGrx1 or

0.03-0.1 μM hGrx2) and preincubated for 5 minutes at 30°C. Then a pre-made complex containing FeCl2 (0.5 mM) and ADP (2.5 mM) was added to each, and the reactions were

initiated by addition of H2O2 (50 μM). Reactions were monitored according to decrease in A340 over 5 minutes (within the linear range of time-dependence). Rates of GSSG formation were determined by measuring time-dependent decrease in A340 (i.e., NADPH

oxidation by GSSG reductase), and enzyme-catalyzed scavenging was calculated by

determining the difference in reaction rates in the presence and absence of Grx, and

expressed as units/mL in the reaction mixture (1 unit = 1 μmol NADPH oxidized/min).

GSH peroxidase activity of hGrx isoforms – GSH-dependent peroxidase activity was

measured via a coupled, spectrophotometric assay identical to that of CSSG

deglutathionylation, except that reactions were initiated by addition of H2O2 (500 µM,

final) instead of CSSG. Enzyme concentrations were as follows: hGrx1, 1 µM; hGrx2, 1

µM; bovine glutathione peroxidase (GPx), 5 nM.

Determination of Grx-mediated protein glutathionylation in the presence of GS•, GSNO,

and GSSG — To ensure that the GAPDH used as a substrate for glutathionylation was

fully reduced, DTT (5 mM, final) was added to stock solutions (~5 mg/mL GAPDH in

0.1 M Na/K-phosphate buffer, pH 7.5) followed by incubation at 4°C for 1 hour. To

remove DTT, the mixture was passed over a DG-10 size exclusion column (Bio-Rad) pre-equilibrated with Na/K-phosphate buffer. Protein-containing fractions (identified by

162 absorbance at 280 nm) were pooled and concentrated to 150-450 μM for use in glutathionylation assays. Glutathionyl radicals were generated by horseradish peroxidase utilizing H2O2 and GSH according to an adaptation of the method of Harman et al.

(1986), who confirmed GS• formation by EPR spectroscopy. Briefly, [35S]-GSH

(0.5mM) and horseradish peroxidase (HRP, 0.2 mg/mL) were included in reaction mixtures (see below) and GS• formation was initiated by addition of H2O2 (50 μM) as described by Starke et al. (2003). The GS• concentration achieved by this radical generating system (i.e., HRP, GSH, and H2O2) was estimated previously (Starke et al.,

2003) to be ≤10 μM based on the amount of GSSG accumulated (GSSG reductase assay); formation of GS-radicals was confirmed by EPR spectroscopy (experiments performed by Dr. Suparna Qanungo (Mieyal Laboratory) in collaboration with Drs. Periannan

Kuppusamy and Mahmood Khan, Ohio State University). For GSNO and GSSG, the radiolabeled GSH derivatives were prepared in advance and used to initiate each reaction.

S-nitrosoglutathione ([35S]-GSNO) was prepared as described by Starke et al. (2003) by

35 combining equimolar amounts of HCl, NaNO2, and [ S]-GSH, and incubating at room temperature for 15 min. Essentially stoichiometric conversion of GSH to GSNO was verified by diluting the stock solution into 15mM HEPES (pH 7.6), conditions in which

-1 -1 ε338 was previously reported for GSNO (980 M cm (Lobysheva et al., 1999)). GSH disulfide ([35S]-GSSG) was prepared by incubating GSSG with trace amounts of [35S]-

GSH for approximately 30 min at room temperature. For kinetics assays, reaction mixtures contained Na/K-phosphate buffer (0.1 M, pH 7.5), rabbit muscle GAPDH (25

μM), Grx (0.02-0.25 μM), and S-carboxymethyl BSA (7 mg/mL) as a co-precipitant to facilitate precipitation of the radiolabeled products. For GS•-mediated glutathionylation,

163

HRP (0.2 mg/mL) and GSH (0.5 mM) were also included. Reactions were initiated with

35 H2O2 (0.05 mM) (for GS•-mediated glutathionylation), [ S]-GSNO (0.05-0.5 mM), or

[35S]-GSSG (0.1-0.3 mM). All reactions took place at room temperature and were

quenched by addition of an equal volume of 20% ice-cold TCA. Precipitated proteins were sedimented by centrifugation at 10,000 xg for 5 min at 4°C, washed, and analyzed for bound radioactivity. Grx-mediated GAPDH-SSG formation was calculated by comparing the bound radioactivity in the presence vs. absence of enzyme. For determination of turnover, rates of protein glutathionylation with and without Grx were determined within the initial, linear phase of the reaction (i.e., first 30 seconds), by dividing nmol protein-SSG formed per second by nmol Grx in each reaction (i.e., turnover number, s-1). All reported turnover values were determined under conditions in which reaction rates exhibited linear dependence upon Grx concentration.

Estimation of concentrations of Grx isoforms in mitochondrial subcompartments – Grx

content (ng/mg total mitochondrial protein) determined by Pai et al. (2007) was divided by the estimated volume of the mitochondrial space per mg mitochondrial protein. The volume of the mitochondrial matrix is calculated by subtracting the mitochondrial volume permeable to radiolabeled sucrose (i.e., intermembrane space and adherent fluid) from that permeable to radiolabeled H2O (i.e., total mitochondrial volume) (Pfaff et al.,

1968). Independent determinations of matrix volume consistently indicate a value ~1.5

μL/mg total protein (Pfaff et al., 1968; Quinlan et al., 1983; Cohen et al., 1985). The volume of the mitochondrial intermembrane space can be determined by subtracting the space permeable to radiolabeled dextran (i.e., the adherent fluid) from the sucrose space

(see above) (Cohen et al., 1985). Measurements of matrix and intermembrane space

164 volumes under a variety of conditions indicated that the intermembrane space volume is consistently ~75% of the matrix volume. Thus, 1.125 μL/mg (75% of 1.5 μL/mg) was used for calculation of Grx1 concentration in the intermembrane space. Molar concentrations of each Grx isoform were determined by dividing the concentration in ng/μL by the molecular weight of hGrx1 (11.7 kDa (i.e., 11,700 ng/nmol)) (Papov et al.,

1994)) in the case of the intermembrane space, or mature hGrx2a (i.e., without the mitochondrial localization sequence, 15 kDa (Lonn et al., 2007)) for the matrix. Then,

Grx(1 or 2) concentration in mg/mL was used to calculate total potential deglutathionylation activity by multiplying by the respective specific activity (U/mg, Table 3.3, p. 185) and by the volume of the appropriate mitochondrial compartment.

Calculation of α-helix 2 dipole moments of hGrx1 and hGrx2: Partial PDB files of each hGrx isoform containing both the catalytic cysteine and helix 2 (hGrx1, 1JHB (Sun et al.,

1998), C22-S33; hGrx2, 2FLS (Johansson et al., 2007), C37-M50), were submitted to the

Protein Dipole Moments Server (http://bioportal.weizmann.ac.il/dipol/) (Felder et al.,

2007) for calculation of the helix 2 dipole moment and helix-thiolate angle (i.e., the angle theta (θ) formed by the intersection of the helix 2 axis with a line projecting from the base of the helix to the catalytic cysteine). By representing the component of the helix dipole directed toward the cysteine thiolate as the base of a right triangle, with the helix axis

(full dipole vector) as the hypotenuse, the magnitude of this component is determined by multiplying the value of the full dipole by cosine θ (adjacent = hypotenuse x cos θ). For hGrx1, we submitted 4 of 20 total NMR structural solutions (structures 1, 6, 16, and 20

(Sun et al., 1998)) shown in a separate study (Jao et al., 2006) to predict a C22 pKa close to the empirical value of 3.5 (Mieyal et al., 1991b; Gan and Wells, 1987). The structure

165 of hGrx2, obtained by x-ray crystallography, contained an associated GSH molecule, which was not included in our calculations.

Statistical analysis – Unless otherwise indicated, data are reported as mean ± standard error (mean ± SEM). Individual data points more than two standard deviations from the mean were considered to be outliers and omitted. Differences between data sets were tested for statistical significance using Student’s t-test (Microsoft Excel) on primary data.

3.4 Results

Characterization of recombinant mammalian Grx2 – Recombinant human and mouse

Grx2 enzymes were purified via a procedure described by Chrestensen et al. (1995) which employs size exclusion chromatography as the primary method of separation of

Grx2 from other E. coli proteins. Figure 3.1A (p. 195) shows a representative elution profile of a G-75 column loaded with E. coli lysate after transformation, induction, collection, lysis, DNA precipitation, and heat treatment (see Materials and Methods).

Fractions exhibiting Grx activity but undetectable protein content (i.e., second protein peak, first activity peak) were collected, concentrated, and used for experiments. By the end of the purification procedure, Grx2 was purified essentially to homogeneity (Figure

3.1B, p. 195). As expected, neither human nor mouse Grx2 reacted with an anti-Grx1 antibody when analyzed by Western blotting (Figure 3.1C, p. 195), while human Grx2 did react with an anti-Grx2 antibody as expected (Figure 3.1D, p. 195, mGrx2 not tested). hGrx2 is selective for glutathione-containing mixed disulfides – To test whether Grx2, like Grx1, is selective for glutathione-containing protein mixed disulfides (i.e., protein-

SSG), we compared the rates of GSH-dependent reduction of two mixed disulfides of

166

bovine serum albumin (BSA) in the absence and presence of Grx2 or Grx1. The first,

BSA-SS-glutathione (BSA-SSG), was specifically prepared to contain a single disulfide-

adducted glutathionyl moiety (see Materials and Methods), and this has been our

prototype protein-SSG substrate for kinetic analysis of glutaredoxin (Gravina and Mieyal,

1993; Srinivasan et al., 1997). The second, BSA-SS-cysteine (BSA-SSC), has the same

disulfide bond to the BSA protein (but not the γ-glutamyl and glycyl moieties of GSH),

yet it does not serve as a substrate for Grx1 (Gravina and Mieyal, 1993; Yang et al.,

1998). Like hGrx1, hGrx2 (Figure 3.2A, p. 197) does catalyze GSH-dependent

dethiolation of BSA-SSG but does not catalyze dethiolation of BSA-SSC (Figure 3.2.B,

p. 197) even when ten-fold excess enzyme is added, documenting high selectivity for

glutathione-containing mixed disulfide substrates. Notably, turnover of BSA-SSG by

Grx2 (39 ± 5 min.-1) was about ten-fold lower than the rate for hGrx1 (457 ± 91 min.-1),

analogous to the ten-fold lower activity of hGrx2 reported previously (Lundberg et al.,

2001) for the pro-substrate hydroxyethyl disulfide (which converts to hydroxyethyl-

SSG).

Grx2 utilizes a nucleophilic, double-displacement (ping-pong) catalytic mechanism – To determine the catalytic mechanism of Grx2, we performed two-substrate kinetic analysis in which the concentrations of CSSG or BSA-SSG were varied at several fixed concentrations of the second substrate GSH, and vice versa; and double reciprocal plots

(v vs. 1/RSSG or 1/GSH) were generated for each fixed concentration of the other substrate. As observed for hGrx1 (Gravina and Mieyal, 1993; Srinivasan et al., 1997), the kinetic analyses of hGrx2 and mGrx2 (Figure 3.3, p. 198) produced parallel line patterns, characteristic of a nucleophilic, double displacement (i.e., ping-pong)

167

mechanism. Secondary plots of 1/Vmax vs. 1/[S] revealed x-intercepts approaching zero

(Figure 3.3, insets, pp. 199-201), indicative of “true KM” values approaching infinity for

each of the substrates. These observations reflect high commitment to catalysis,

precluding reversible enzyme-substrate complexes in the catalytic mechanism,

corresponding to the kinetic behavior characteristic of hGrx1 (Gravina and Mieyal,

1993). Additional support for this essentially irreversible encounter-type mechanism was

the observation that neither hGrx2 or mGrx2 was inhibited by the GSH analog S- methylglutathione at concentrations up to 3mM (i.e., six-fold excess of GSH (Table 3.1,

p. 185), a behavior also reported for hGrx1 (Srinivasan et al., 1997).

Grx2 exhibits decreased catalytic efficiency towards cysteine-SSG – Previously, hGrx2 was reported to exhibit ~5-fold lower apparent KM values for multi-glutathionylated BSA and RNase substrates relative to hGrx1, leading to the proposal that Grx2 exhibits high affinity for glutathionylated proteins (Johansson et al., 2004). On the contrary, 1/v vs.

1/S curves from double-reciprocal plots (above, Figure 3.3, pp. 199-201) indicated that apparent KM values for the monoglutathionylated BSA-SSG substrate were within a factor of 2 for both Grx isoforms (Table 3.2.A, p. 187). On the other hand, a marked decrease in apparent kcat of ~10-fold (Grx2 < Grx1) was observed when either BSA-SSG

or GSH was used as the substrate whose concentration was varied. This diminution in

kcat corresponds to a diminished catalytic efficiency for Grx2 (human and mouse) relative

to Grx1 (Table 3.2, p. 187).

The discrepancy between KM,app values in this study compared to a previous report

(Johansson et al., 2004) is not clear; however, it is conceivable that the observed differences in KM,app reflect differences in the glutathionylated protein substrates used by

168

each group. To eliminate steric considerations contributed by glutathionylated protein

substrates, we determined apparent kinetic constants for each Grx isoform using the

prototype substrate cysteine-SSG (CSSG). An additional advantage to the CSSG

substrate is that it represents the conserved moiety among all protein-cysteinyl-SS-

glutathione (protein-SSG) substrates, making it the logical focus of “high-affinity” of

protein-SSG for Grx2, if this is an intrinsic property of the enzyme. Under standard

assay conditions (see Materials and Methods), human and mouse Grx2 both exhibited

about 9-fold lower kcat values for CSSG compared to hGrx1, while apparent KM values differed by only 1.2-fold, resulting in a decreased catalytic efficiency of Grx2 similar to that observed for BSA-SSG (Table 3.2.C, p. 187). Thus, for all substrates tested, the ratio of kcat,app to KM,app (i.e., catalytic efficiency) was decreased by nearly 10-fold for

human and mouse Grx2 compared to Grx1, with the decrease driven primarily by

diminished apparent kcat.

Grx2-mediated deglutathionylation operates via a monothiol mechanism – Mammalian thiol-disulfide oxidoreductase enzymes utilize distinct catalytic mechanisms, as exemplified by Grx1 (i.e., monothiol mechanism) and Trx (i.e., dithiol mechanism), as

illustrated in Scheme 3.1B, p. 184, (reviewed in Mieyal et al. (1995)). To determine

which mechanism is utilized by Grx2, we compared the activity of wild type enzyme to

one in which the second cysteine at the active site was mutated to serine (i.e., C40S).

Like the analogous hGrx1 mutant (i.e., C25S), hGrx2-C40S exhibited increased specific

activity compared to wild type hGrx2 (Table 3.3, p. 188), suggesting that C40 is not

required for catalysis, and it actually detracts from the catalytic rate. Under standard

assay conditions, hGrx2 C40S exhibits a lower KM,app for GSH (Figure 3.4A, p. 203)

169

compared to wild type enzyme, and approximately 2-fold increases in kcat,app and KM,app

for CSSG (Figure 3.4B, p. 203). These results suggest that the C-terminal cysteine at the

active site of Grx2, like that of hGrx1 (Yang et al., 1998), competes with GSH for

nucleophilic attack on the hGrx2-SSG intermediate and forms the hGrx2-intramolecular

disulfide as shown in Scheme 3.1A, p. 184. Notably, the magnitude of the increases in

kcat and KM for CSSG exhibited by hGrx2-C40S is similar to that observed for the analogous hGrx1 mutant (Grx1-C25S (Yang et al., 1998)), suggesting the relative steady-

state concentrations of the Grx-SSG and intramolecular disulfide forms are approximately the same for the two Grx isoenzymes. pH rate profile of hGrx2-dependent deglutathionylation - The pH rate profile of hGrx1- catalyzed deglutathionylation of BSA-SSG displays a typical sigmoid titration curve with an inflection point near pH 8.5, corresponding to the pKa of GSH and indicating the rate- determining step of catalysis as nucleophilic attack of the GS-thiolate on the hGrx1-SSG intermediate (i.e., Scheme 3.1A, p. 184, Step 2 (Srinivasan et al., 1997)). Likewise, the pH-dependence of hGrx2-mediated deglutathionylation of BSA-SSG also exhibits an inflection point near 8.5 (Figure 3.5, p. 204), superimposable on the pH rate profile of the non-enzymatic reaction of GSH with BSA-SSG (Figure 3.5, p. 204 inset), indicating

the limiting factor in both reactions is the titration of the GSH-thiol, as observed

previously for Grx1 (Srinivasan et al., 1997). Thus the Grx2-mediated reaction displays

the same rate-determining step (Scheme 3.1A, p. 184, Step 2,). This observation differs

from previous reports of maximal activity at pH 8.5 (i.e., a bell-shaped curve) for rat liver

Grx1 (Gan and Wells, 1986), and for human Grx1 and Grx2 (Johansson et al., 2004).

This discrepancy likely relates to an artifact of the assay for Grx activity in these previous

170

reports. In the GR-coupled assay an apparent maximum near pH 8.5 is observed because

of the pH sensitivity of GR activity. If enough GR is added at higher pH, a typical

single-component titration curve is achieved (as described for Grx1 in Gravina and

Mieyal, 1993). To avoid this artifact we used the radiolabel assay which monitors

formation of GSSG in the absence of GR.

The pKa of Grx2’s catalytic Cys is 4.6 – For typical thiol-disulfide exchange reactions, each one pH unit decrease in pKa of the leaving group thiol predicts a ~4-fold increase in

second-order rate constant (Szajewski and Whitesides, 1980). For Grx1-catalyzed

protein deglutathionylation, the rate-determining step is thiol-disulfide exchange between the Grx1-SSG intermediate and GSH (Srinivasan et al., 1997), and the difference in pKa

between the leaving group in that reaction (i.e., ~3.5 for Grx1-SH) and the leaving group

of the uncatalyzed reaction (i.e., ~8.5 for a typical protein-SH) accounts for the majority

of the rate enhancement of Grx1-mediated deglutathionylation (4ΔpKa = 45 = 1024-fold

(Srinivasan et al., 1997). Here, we have shown that Grx2 operates via an analogous ping- pong mechanism (Figure 3.3, pp. 199-201), in which attack of GS- on the Grx2-SSG intermediate is also rate-determining (Figure 3.5, p. 204). To investigate whether the diminished kcat of hGrx2 towards glutathionylated substrates could be explained by a

higher pKa of its catalytic cysteine, we determined its pKa according to the pH

dependence of hGrx2 inactivation by iodoacetamide (IAM). hGrx2’s catalytic Cys was

found to have a pKa of 4.6 (Figure 3.6, p. 205), approximately 1 pH unit higher than that

of Grx1, accounting for about half of the ~10-fold decrease in specific activity (see

Discussion).

171

Contribution of Grx helix 2 dipole moment to stabilization of the catalytic cysteine

thiolate – The basis for the low pKa of hGrx1’s catalytic cysteine is uncertain, but

interaction with the dipole of helix 2 has been suggested to contribute (Jao et al., 2006;

Kortemme and Creighton, 1995). To determine whether a decreased ion-dipole interaction between helix 2 and the catalytic cysteine thiolate explained the higher pKa of

hGrx2, we used published structures of hGrx1 and hGrx2, in combination with a dipole

calculation program (http://bioportal.weizmann.ac.il/dipol/ (Felder et al., 2007)), to

determine the component of the helix 2 dipole moment directed toward each enzyme’s

catalytic cysteine (see Materials and Methods). Surprisingly, the component of the helix

2 dipole was greater for hGrx2 than for each of the hGrx1 structures analyzed (Table 3.4,

p. 189) predicting greater stabilization of its catalytic cysteine thiolate by the helix.

Therefore, the increased pKa of hGrx2 cannot be explained by a decreased ion-dipole

interaction with helix 2, and is likely explained by other electrostatic interactions (see

Discussion).

Grx2 exhibits decreased enhancement of the nucleophilicity of GSH compared to Grx1 –

Rate enhancement of protein deglutathionylation by Grx1 is attributed both to the low

pKa of its catalytic cysteine (see above) and to its ability to enhance the nucleophilicity of the second substrate, GSH, beyond that predicted by its pKa, as shown by Bronsted

analysis (Srinivasan et al., 1997). The preference for GSH as the second substrate

appears to be due mainly to the γ-glutamylcysteine moiety as demonstrated in previous studies of hGrx1 (Srinivasan et al., 1997) and E. coli Grx1 (Peltoniemi et al., 2006a). To

determine whether hGrx2, like hGrx1, enhances the nucleophilicity of GSH, we

determined the rate enhancement of hGrx1 and hGrx2-mediated deglutathionylation of

172

BSA-SSG using GSH, and the closely related cysteinylglycine, whose nucleophilicity is

not enhanced by human Grx1 (Srinivasan et al., 1997). Indeed, second-order rate

constants for deglutathionylation by cysteinylglycine reflected only the rate enhancement

predicted by the pKa of each Grx enzyme’s catalytic cysteine (Table 3.5, p. 191, and

Discussion). When cysteinylglycine was replaced by GSH, rate enhancement was

augmented by ~20-fold for hGrx1, but by less than 10-fold for hGrx2 (Table 3.5, p. 191).

Thus, hGrx1 appears to enhance GSH’s ability to serve as a second substrate to a greater

degree than does hGrx2.

The GSH/GR system is the preferred coupling system for Grx1 and Grx2 even under

“oxidative stress” conditions – In the absence of GSH, hGrx2-mediated reduction of

CSSG was shown to be augmented somewhat by NADPH and thioredoxin reductase

(TR) (Johansson et al., 2004). Under GSH-free conditions, we observed TR-facilitated

CSSG reduction by both Grx1 and Grx2, displaying similar rates for the two isoforms, comparable to the rate reported previously for hGrx2 (Johansson et al., 2004).

Remarkably, replacement of TR with GSH and GSSG reductase (GR)—at concentrations

10-fold lower than found in cells (Carlberg and Mannervik, 1985)—increased CSSG

deglutathionylation rates by 20-fold for Grx2, and >200-fold for Grx1 (Table 3.6, p.

192). Addition of TR (with or without Trx) to the reaction mixtures containing GSH and

GR did not further augment CSSG turnover in either case. Therefore, while TR can

assist deglutathionylation by both Grx isoforms to some degree, it is substantially less

efficient than the GSH/GR system, even under conditions of depleted GSH, as might

occur under oxidative stress conditions in vivo (see Discussion).

173

Like Grx1, Grx2 exhibits glutathione-thiyl radical scavenging activity – Under conditions of glutathione-thiyl radical (GS•) generation, Grx1 catalyzes GSSG formation (Starke et al., 2003). Likewise, in the current study we observed that hGrx2 catalyzed GSSG formation when GS• was produced; however turnover was about 10-fold lower than that of hGrx1 (Figure 3.7A, p. 207). As shown previously for hGrx1 (Starke et al., 2003), hGrx2-mediated formation of GSSG under GS-radical generating conditions could not be explained by the enzyme acting as a GSH peroxidase (Figure 3.7B, p. 207).

Like Grx1, Grx2 can promote protein S-glutathionylation in the presence of various activated glutathionyl donors – Previously, we demonstrated that hGrx1 promotes protein

S-glutathionylation of model protein substrates in the presence of GS•, GSNO, or GSSG; with GS• being the most efficient GS-donor (Starke et al., 2003). Here we compared rates of glutathionylation of the model protein GAPDH by hGrx1 and hGrx2. For all substrates tested, the majority of GAPDH glutathionylation occurred within the initial, linear phase of the reaction time course≤ ( 30s, data not shown). Like hGrx1, hGrx2 promoted GAPDH-SSG formation in the presence of all three GS-donors, with GS• being a far superior substrate than GSNO and GSSG (Table 3.7, p. 193). With GSSG, hGrx1 was a better catalyst of GAPDH glutathionylation than Grx2, i.e., ~4-fold higher turnover, characteristic of the active site pKa difference (see Discussion). However, equivalent rates of GAPDH glutathionylation for the two Grx isoforms were observed with GS• and GSNO.

3.5 Discussion

Kinetic comparison of mammalian Grx isoforms – Earlier reports have emphasized differences between the human Grx isoforms (Johansson et al., 2004; Fernando et al.,

174

2006). In contrast, we have documented remarkable catalytic similarities between Grx2

and Grx1, including a double-displacement deglutathionylation mechanism (Scheme

3.1A, p. 184) with the same rate determining step, which cycles through an intermediate involving only the N-terminal active site cysteine residue (monothiol mechanism), and exhibits high commitment to catalysis. An important consequence of this mechanism in vivo is that kcat and KM values for protein-SSG substrates increase or decrease in parallel

with the concentration of GSH; therefore, maximal rates of deglutathionylation by both

enzymes will depend upon the redox balance of the local environment. Another

similarity between the Grx enzymes is that the GSH/GR system is superior to the Trx

system in supporting deglutathionylation, even when GSH and GR were diminished to

one-tenth of their predicted intracellular concentrations. Finally, hGrx2 (like hGrx1)

exhibited GS• scavenging activity, as well as catalysis of protein-SSG formation, with

GS• acting as the best activated glutathionyl co-substrate.

A striking distinction between mammalian Grx1 and Grx2 is their difference in

catalytic efficiency, reflecting ~10-fold lower kcat values for hGrx2 towards CSSG, BSA-

SSG, and GSH, accompanied by only minor changes in KM,app. We used our

understanding of the basis for Grx1-mediated rate enhancement to investigate why Grx2

exhibits a lower kcat, and found that the ~10-fold difference in kcat between hGrx1 and h/mGrx2 can be explained by two factors: (1) a 1 pH unit increase in pKa of Grx2’s

catalytic cysteine, which predicts a ~4-fold lesser rate enhancement vs. hGrx1, (i.e., 4ΔpKa

= 44, or 256-fold vs. 1024-fold), and (2) a diminished enhancement of nucleophilicity of

GSH.

175

For Grx1, the basis for enhancement of the nucleophilicity of GSH, as well as for the

low pKa of its catalytic cysteine, are not fully understood; however, structural and site-

directed mutagenesis studies provide some insight. For example, a recent analysis of the

role of K19 in rate enhancement by hGrx1 (Jao et al., 2006) suggested that the primary

role of this positively charged moiety is not to stabilize the catalytic cysteine thiolate

anion (as was inferred from its close proximity to C22 in the average NMR structure (Sun

et al., 1998)), but rather to enhance deglutathionylation by another mechanism, such as enhancing the nucleophilicity of GSH. Interestingly, structural comparison of reduced hGrx1 (Sun et al., 1998) and hGrx2 (Johansson et al., 2007) indicates a substantial shift

in orientation of hGrx2’s K34 (homologous to K19 of hGrx1) to a position further from

the catalytic cysteine (~3Å for hGrx1 vs. 8.2Å for hGrx2). The effect of this shift in orientation on enzyme-substrate interactions is unknown, but could be explored by determining the effect of K34 mutation on rate enhancement (and pKa) of hGrx2.

Several hypotheses have been proposed to explain the low pKa of Grx1’s catalytic

cysteine (C22) besides proximity of positively charged residues, including ion-dipole

interactions between C22 and helix 2 (Jao et al., 2006; Kortemme and Creighton, 1995),

H bonding between the C22-thiolate and C25 and/or T21 side chains (Sun et al., 1998;

Jao et al., 2006), and H bonds between the C22-thiolate and backbone residues within the active site (Foloppe and Nilsson, 2007). Calculation of the helix 2 dipole moment for each hGrx isoform revealed that the dipole vector component was greater for Grx2 than for hGrx1, indicating that a diminished helix 2 dipole does not explain the higher pKa of

the Grx2 catalytic thiolate.

176

Increased H bonding distances within the hGrx2 active site also do not explain its

higher pKa relative to hGrx1. Foloppe and Nilsson (2007) proposed that for pig Grx1

(pGrx), H bonds between the catalytic cysteine thiolate and amide hydrogens of active site residues P23 and C25 account for the low catalytic cysteine pKa (~3.5). However, calculated distances between Grx2’s C37 and homologous Y39 and C40 amide nitrogens are all smaller than the distances between hGrx1’s C22, Y24, and C25 backbone nitrogens. In fact, the only interaction proposed to stabilize the hGrx1 C22-thiolate that appears to be diminished for hGrx2 is the H bond between the catalytic cysteine thiolate and the hydroxyl group of T21, which is supported by only 1 of 20 hGrx1 NMR structures (2.2 Å for hGrx1, structure 16; 9.4Å for hGrx2). Importantly, these comparisons are complicated by the fact that the protein structures were solved using different methodologies (i.e. NMR vs. X-ray crystallography), and that the structure of hGrx2 contains an associated GSH that may alter active site orientation. Determination of the structure of reduced hGrx2 will facilitate more direct structural comparison, providing additional insight into the basis for the unique pKas of hGrx1 and hGrx2.

Additional activities of Grx2; Implications of GS-transfer activities – hGrx1 was

previously shown to scavenge glutathione thiyl radicals (GS•), likely via initial

stabilization of the radical by formation of a Grx-SSG•- disulfide anion radical

intermediate (Scheme 1.2C, p. 112 (Starke et al., 2003)). Rate enhancement by hGrx1

was explained by the low pKa of its catalytic cysteine (which serves as a nucleophile in

the first step of the reaction, and is >99% ionized at physiological pH), as well as its

ability to stabilize a covalently bound glutathionyl moiety (Yang et al., 1998). We

hypothesized that hGrx2 would also exhibit GS-scavenging activity, based upon its low

177 pKa (4.6, > 99% ionized at physiological pH) and conserved residues implicated in stabilization of the adducted glutathionyl moiety (Gladyshev et al., 2001; Lundberg et al.,

2001). The 10-fold lower rate of GS• scavenging by hGrx2 compared to hGrx1 suggests that the final step of the reaction (i.e., Scheme 1.2C, p. 112, step 3)—which is identical to the rate-limiting step of deglutathionylation (i.e., Scheme 3.1A, p. 184, step 2)—is also rate-determining for GS• scavenging. As shown previously for hGrx1 (33), hGrx2- mediated GS• scavenging can not be explained by the enzyme acting as a GSH peroxidase. Although GSH-peroxidase activities were reported recently for hGrx2

(Fernando et al., 2006), >5 μM Grx2 was required for detection of activity, a concentration more than 100 times what was effective in GS• scavenging, and more than

5 times the predicted Grx2 concentration within the mitochondrial matrix (see below).

Mechanisms of protein S-glutathionylation in vivo are not yet elucidated; however, involvement of oxidized derivatives of GSH has been proposed (Klatt and Lamas, 2000;

Shelton et al., 2005; Gallogly and Mieyal, 2007; Ghezzi, 2005b; Hurd et al., 2005), and

Starke et al. (Starke et al., 2003) provided evidence that such reactions can be mediated by hGrx1 in vitro. Like hGrx1, hGrx2 promoted transfer of a glutathionyl moiety to

GAPDH in the presence of various oxidized derivatives of GSH, with GS• acting as a far superior substrate than GSNO and GSSG. For GSSG, hGrx2 exhibited about one-fourth the glutathionylation rate mediated by hGrx1. This observation suggests that the protein glutathionylation reaction with GSSG proceeds via a thiol-disulfide exchange mechanism analogous to the deglutathionylation reaction (i.e., Scheme 3.1A, p. 184, with Protein-

SSG replaced by GSSG, and GSH replaced by Protein-SH, as documented previously for hGrx1 (Mieyal et al., 1991b), and discussed in (Mieyal et al., 1995); i.e., the reverse

178

reaction. The decreased activity of hGrx2 in this case is accounted for entirely by the

difference in pKa of the catalytic cysteine, because there is no enhancement of

nucleophilicity for the GAPDH-cysteine-thiol moiety by either enzyme. Importantly, the

GSSG concentration required to achieve turnover by this mechanism (≥ 0.1 mM)

suggests that it is unlikely to contribute significantly to protein glutathionylation under

redox signaling conditions in vivo, in which the GSH:GSSG ratio does not deviate

substantially from the typical value of 100, corresponding to micromolar concentrations

of GSSG for most cells.

For GS• and GSNO, hGrx2 promoted GAPDH glutathionylation at rates equivalent to

those of hGrx1, representing a remarkable contrast to its lower deglutathionylation and

GS• scavenging activities. These equivalent catalytic rates suggest that the rate-

determining step of Grx-mediated protein-SSG formation—in contrast to catalysis of

deglutahionylation—does not involve the catalytic cysteine serving as a leaving group.

This concept is consistent with the mechanism proposed previously for GS•-transfer by

hGrx1 (Scheme 1.2D, p. 112 (Starke et al., 2003)). For both Grx enzymes, GS• was a much better glutathionyl-donor substrate compared to GSNO and GSSG, and thus represents a more likely substrate for protein S-glutathionylation by Grx in the absence of overt oxidative or nitrosative stress. Interestingly, while GSNO may not serve as a direct substrate for Grx-mediated glutathionylation under resting conditions, it may still contribute to protein-SSG formation by reacting with GSH to form the superior substrate,

GS• (Singh et al., 1996). The possibility of GSNO dismutation, combined with the observation that a ten-fold molar excess of GSNO disrupted inactive Grx2 dimers in vitro

(Hashemy et al., 2007), suggests a potential role for GSNO as an activator of Grx2-

179

mediated glutathionylation in vivo, first by releasing the active enzyme from its inactive form, and then by reacting with GSH to generate a more efficient substrate for protein

glutathionylation.

In vivo implications of Grx2 activities – A major distinction between mammalian Grx

isoforms is their nearly 10-fold difference in specific activity for deglutathionylation of

glutathionyl mixed-disulfide substrates. However, deglutathionylation activity in vivo is

likely to be determined by multiple additional considerations, such as intracellular

enzyme concentrations, pH (~8 in the mitochondrial matrix (Llopis et al., 1998)), and

activating or inactivating factors. We estimated the intramitochondrial concentrations of

Grx1 and Grx2a using the results of our laboratory’s previous study of Grx content in rat

mitochondria (i.e., ng Grx/mg total mitochondrial protein) (Pai et al., 2007) in combination with the specific volume estimated for each of the intramitochondrial compartments (i.e., μL/ mg mitochondrial protein) (Pfaff et al., 1968; Cohen et al., 1987)

(see Materials and Methods). Accordingly, the concentration of Grx2 in the

mitochondrial matrix is estimated to be ~1 μM, while that of Grx1 in the intermembrane

space is about 10-fold lower (~0.1 μM). Assuming that both Grx isoforms are fully

active, the ten-fold higher Grx2 concentration—balanced by its 10-fold lower kcat—

predicts that total deglutathionylation and GS• scavenging activities should be similar

across mitochondrial subcompartments. In contrast, Grx-mediated glutathionyl-transfer

activity, catalyzed at equivalent rates by both Grx isoforms, would be expected to be

higher in the mitochondrial matrix, where Grx2 concentration is ten-fold greater.

Prediction of Grx2’s intracellular activity is further complicated by the

observation that the protein may be sequestered into inactive FeS cluster dimers,

180

according to immunoprecipitation studies performed under resting conditions in cultured

cells (Lillig et al., 2005). Regulation of dimer integrity in vivo is not yet understood, but

exposure to oxidants (e.g., GSSG, GSNO) in vitro results in dimer dissociation (Lillig et

al., 2005; Hashemy et al., 2007). The concept that Grx2 may be released (and indeed, activated) primarily under oxidative conditions suggests that its catalytic activities involving oxidized substrates (e.g., GS• scavenging, glutathionyl transfer) may represent its primary functions within the mitochondria. Alternatively, Grx2 may be released upon oxidative stress, but primarily catalyze deglutathionylation, reestablishing sulfhydryl homeostasis, before being re-sequestered into inactive dimers. Clearly more studies are necessary to unravel the roles of Grx2 in situ.

Knock-out and overexpression studies of hGrx2 in HeLa cells (Lillig et al., 2004;

Enoksson et al., 2005) and transgenic mice (Nagy et al., 2008) suggest that Grx2 confers

pro-survival effects during oxidative stress; but the mechanisms of these protective

effects are unknown. Our studies suggest that Grx2 could promote protein

glutathionylation or deglutathionylation depending upon the redox status of the local

environment, and the presence of specific oxidized derivatives of GSH. The distinct

effects of glutathionylation on mitochondrial respiratory chain components suggest that

either activity could be cytoprotective. For example, glutathionylation of Complex II in

- vitro increases its electron transfer efficiency and decreases O2∙ production, a trigger of

apoptosis in disease states such as cardiac ischemia-reperfusion (Chen et al., 2007).

Conversely, deglutathionylation of Complex I restores its electron transfer activity,

- preventing electron leakage and O2∙ production (Beer et al., 2004; Taylor et al., 2003).

Interestingly, isolated hGrx2 (> 6 μM) was shown recently to promote glutathionylation

181

of the 75 kDa subunit of Complex I under oxidizing conditions (i.e., GSH:GSSG = 3) in vitro (Beer et al., 2004); however, the physiological relevance of this observation is yet to be determined. Further studies focused on the relationship between Grx2 activity and glutathionylation status of established targets, or as yet unknown mitochondrial targets, will increase understanding of the mechanism by which Grx2 confers its cytoprotective effects.

In summary, we found that mammalian Grx2 exhibits remarkably similar catalytic properties to hGrx1 (e.g., monothiol, nucleophilic double-displacement catalytic mechanism, high commitment to catalysis, selectivity for protein-SSG substrates, GS• scavenging and glutathionyl transfer activities), as well as some unique features, such as a higher catalytic cysteine pKa and decreased enhancement of GSH nucleophilicity, resulting in decreased kcat,app for both substrates in deglutathionylation reactions. The

major catalytic function of Grx2 in vivo remains to be characterized, but this pursuit

represents an exciting frontier in the fields of redox signaling, mitochondrial redox

homeostasis, and cellular viability.

182

Scheme 3.1. Catalytic mechanisms of mammalian glutaredoxin (Grx) and

thioredoxin (Trx). A, Catalytic mechanism of protein deglutathionylation by mammalian Grx1 (Gravina and Mieyal, 1993; Yang et al., 1998) and Grx2. In the first

step, the catalytic cysteine-thiolate of Grx attacks the glutathionyl sulfur of the protein-

SSG mixed disulfide, releasing the first product, protein-SH. In the second step of the reaction, GSH attacks the enzyme-SSG intermediate, restoring the reduced enzyme and

releasing GSSG. Alternatively, the cysteine adjacent to the active site may attack the

Grx-SSG intermediate, forming an intramolecular disulfide and releasing GSH. The

intramolecular disulfide may be recruited back into the catalytic cycle by reaction with

GSH, reforming Grx-SSG, and proceeding with step 2. Reduction of GSSG is catalyzed

by GSSG reductase, coupled to oxidation of NADPH. PSSG, protein-GSH mixed

disulfide; Grx, glutaredoxin; GR, GSSG reductase. B, Catalytic mechanism of

intramolecular disulfide reduction by mammalian Trx (Holmgren, 1985). In the first step

of the reaction, the acidic cysteine-thiolate of Trx attacks an intramolecular protein

disulfide, forming a transient HS-Trx-SS-protein-SH intermediate in which the second active-site cysteine of Trx attacks the intermolecular disulfide and displaces protein-

(SH)2,. forming Trx-(S)2 intramolecular disulfide. Trx disulfide is then reduced by Trx reductase (TR), coupled to oxidation of NADPH. P, protein; Trx, thioredoxin; TR, Trx

reductase.

183

184

Table 3.1. S-methylglutathione does not inhibit Grx2-catalyzed

deglutathionylation. Activities of hGrx1 (8 nM), hGrx2 (37 nM), and mGrx2 (80 nM)

were analyzed via the spectrophotometric assay (see Materials and Methods) in the presence and absence of 3 mM S-methylglutathione (GSMe). Percent inhibition was calculated by subtracting the rate in the presence of inhibitor from the rate in the absence of inhibitor, then dividing by the rate in the absence of inhibitor and multiplying by

100%. Comparison of reaction rates in the absence of Grx indicated that GSMe inhibits

GR, the coupling enzyme in the spectrophotometric assay, by 18%. Since there was no additional inhibition of the reaction in the presence of Grx, it was concluded that GR inhibition accounted for the decreased reaction rates in reactions containing Grx and

GSMe. Data represent mean % inhibition calculated from paired experiments (± GSMe)

± standard error. For reactions without Grx, n = 18; for hGrx1, n = 5; for hGrx2, n = 6; for mGrx2, n = 5.

p-value Grx added % inhibition vs. -Grx vs. Grx1 none 18.3 ± 1.8 hGrx1 15.3 ± 2.8 0.4 hGrx2 16.3 ± 3.4 0.6 0.8 mGrx2 11.6 ± 2.7 0.1 0.4

185

Table 3.2. Catalytic efficiencies of Grx isoforms. Rates of deglutathionylation by recombinant hGrx1, hGrx2 and mGrx2 were determined using the spectrophotometric coupled assay (see Materials and Methods). A, BSA-SSG was varied (0.005-0.25mM), and GSH was maintained at 0.5mM; B, GSH (0.1-1mM) was varied and CSSG was maintained at 0.1mM; C, CSSG was varied (0.002-0.5mM) and GSH was maintained at

(0.5mM). Three separate experiments were performed to generate plots of v (min.-1) vs.

[S]. These data were fit to the Michaelis-Menten equation, and apparent kcat and KM values were calculated accordingly. ND, not determined. Data for BSA-SSG generated by David Starke.

186

A. BSA-SSG -1 -1 -1 Grx Isoform kcat (min. ) KM (mM) kcat/KM (min. mM ) hGrx1 513 ± 19 0.034 ± 0.004 1.5 ± 2.6 × 104 hGrx2 46 ± 2 0.016 ± 0.003 2.8 ± 0.6 × 103 mGrx2 ND ND ND

B. GSH -1 -1 -1 Grx Isoform kcat (min. ) KM (mM) kcat/KM (min. mM ) hGrx1 4.8 ± 0.23 × 103 0.81 ± 0.1 6.0 ± 1 × 103 hGrx2 675 ± 71 1.6 ± 0.4 433 ± 145 mGrx2 804 ± 81 1.3 ± 0.3 614 ± 203

C. CSSG -1 -1 -1 Grx Isoform kcat (min. ) KM (mM) kcat/KM (min. mM ) hGrx1 2.3 ± 0.15 × 103 0.055 ± 0.01 4.2 ± 1 × 104 hGrx2 217 ± 16 0.036 ± 0.009 6.1 ± 2 × 103 mGrx2 286 ± 17 0.055 ± 0.01 5.2 ± 1.3 × 103

187

Table 3.3. Specific activities of Grx isoforms for model substrate Cysteine-SSG

(CSSG) – Rates of CSSG deglutathionylation by recombinant human Grx1 (hGrx1) and

truncated human and mouse Grx2 (hGrx2, mGrx2) were determined using the

spectrophotometric coupled assay described previously (Chrestensen et al., 2000;

Srinivasan et al., 1997; Jao et al., 2006). Reaction mixtures contained sodium/potassium

phosphate buffer (0.1 M, pH 7.4), NADPH (0.2 mM), GSH (0.5 mM), GSSG reductase

(2 U/mL), and Grx (15 nM Grx1, 64-75 nM Grx2), and were initiated by addition of

CSSG (0.1 mM, final) and monitored over 5 min. Rates of deglutathionylation were

determined by measuring the rate of NADPH-dependent reduction of the reaction product

GSSG and subtracting the non-enzymatic rate. Data are expressed as the mean +/-

standard error (n = 4-12).

Grx Isoform Specific Activity (U/mg) Ratio (Grx1:Grx2) hGrx1 106.8 ± 2.8 - hGrx2 11.2 ± 0.05 9.5 mGrx2 12.6 ± 0.9 8.5 hGrx2 (C40S) 20.9 ± 1.4 -

188

Table 3.4. Helix 2 dipole moments for hGrx1 and hGrx2 – Partial pdb files of hGrx1

(1JHB (Sun et al., 1998)) and hGrx2 (2FLS (Johansson et al., 2007)) were submitted to

the Protein Dipole Moments Server (http://bioportal.weizmann.ac.il/dipol/, (Felder et al.,

2007) to calculate the dipole moment of helix 2, as well as the angle between the helix

and catalytic cysteine (see Materials and Methods). For hGrx1, the number in

parentheses indicates the specific NMR structure shown previously to predict the

empirically observed catalytic cysteine pKa of approximately 3.5 (Jao et al., 2006). The

component of each dipole directed toward the catalytic cysteine was calculated by

multiplying the dipole moment by the cosine of the angle formed by the intersection of

the helix axis with a line projecting from the base of the helix to the catalytic cysteine.

Grx Dipole moment Helix-thiolate angle Component of structure (D) (°) dipole (D) hGrx1 (1) 182 24.4 166 hGrx1 (6) 170 19.7 160 hGrx1 (16) 177 19.2 167 hGrx1 (20) 168 24.4 153 hGrx2 213 10.6 209

189

Table 3.5. Enhancement of GSH nucleophilicity by Grx isoforms. Time-dependent release of [3H]-GSSG from [3H]-BSA-SSG was measured as described in Materials and

Methods. Reaction mixtures contained 0.1M AMPSO buffer (pH 9.5), GSH or cysteinyl glycine (Cys-Gly) (0.25 mM, final), and Grx (3-9 nM). Enzyme and substrate

concentrations were within the range of linear dependence (determined in separate

experiments). Second order rate constants (M-1 min.-1) for deglutathionylation were calculated as described by Srinivasan et al. (1997), i.e., rate of GSSG release = k[BSA-

SSG][RSH] (for the uncatalyzed reaction) or k[Grx-SSG][RSH] (for the catalyzed

reaction). Final [GSH] and [Cys-Gly] concentrations at pH 7.5 were calculated according to their pKas (GSH = 8.7, Cys-Gly = 8.8, (Srinivasan et al., 1997)). Rate constants

represent averages of 3-7 determinations ± standard error.

190

191

Table 3.6. Contribution of TR to Grx-mediated deglutathionylation of CSSG. Rates of deglutathionylation of CSSG were measured using the spectrophotometric assay (see

Materials and Methods). Reaction mixtures contained sodium/potassium phosphate buffer (0.1 M, pH 7.5), NADPH (0.2 mM), and hGrx1 (0.37 μM in experiments containing GSH, 3.7 μM in experiments without GSH) or hGrx2 (3.7 μM). When present, other components included GSH (0.1mM), GSSG reductase (0.01 μM), Trx (10

μM), TR (230 nM). Reactions were initiated by addition of CSSG (0.001-0.5 mM, final) and monitored over 5 min. Turnovers represent the average of at least 3 experiments and are expressed as the mean ± standard error.

CSSG turnover by CSSG turnover by Reducing System hGrx1 (min. -1) hGrx2 (min. -1) GSH, GR 196.9 ± 14.5 12.98 ± 0.69 TR 0.94 ± 0.13 0.62 ± 0.08

192

Table 3.7. hGrx1 and hGrx2 exhibit equal rates of protein glutathionylation in the

presence of GS• and GSNO but not GSSG. Reaction mixtures contained

sodium/potassium phosphate buffer (0.1 M, pH 7.5), rabbit muscle GAPDH (25 μM),

Grx (0.02-0.25 μM), S-carboxymethyl BSA (7 mg/mL), and [35S]-GSNO (500 μM),

[35S]-GSSG (500 μM) or GSH (0.5 mM) and GS• (approximately 10 μM, see Starke et

al., 2003). Reactions took place at room temperature, and were quenched by addition of

20% ice-cold TCA. Precipitated proteins were pelleted by centrifugation, washed, and

analyzed for bound radioactivity. Grx-mediated GAPDH-SSG formation was calculated

by comparing the bound radioactivity in the presence vs. absence of enzyme. Rates are

expressed as nmol protein-SSG/min. per nmol Grx (i.e., turnover, min.-1). Turnovers

were determined within the linear range of Grx dependence for each substrate, and are

expressed as the mean +/- standard error (n = 3-12).

Protein-SSG formation Substrate Grx1-mediated (s-1) Grx2-mediated (s-1) GS• (≤10 μM, 0.43 ± 0.03 0.38 ± 0.04 (Starke et al., 2003)) GSNO (500 μM) 0.21 ± 0.07 0.24 ± 0.08 GSNO (250 μM) 0.09 ± 0.04 0.11 ± 0.07 GSSG (250 μM) 0.38 ± 0.01 0.12 ± 0.007 GSSG (100 μM) 0.24 ± 0.03 0.06 ± 0.003

193

Figure 3.1. Characterization of recombinant mammalian Grx2. A, Protein concentration and Grx activity in fractions eluted from a G-75 size exclusion column. E. coli bacteria induced to express recombinant human or mouse Grx2 were collected, lysed, and processed as described in Materials and Methods. Following heat treatment and centrifugation, the supernatant was loaded on a G-75 size-exclusion column pre-

- equilibrated with NH4HCOO (pH 5.8) and fractions were analyzed for protein content

(BCA Assay, Pierce) and Grx activity (spectrophotometric assay, see Materials and

Methods). Fractions corresponding to the highest Grx activity were pooled and concentrated, glycerol was added (20% final), and aliquots were stored at -80 °C. B,

Coomassie-stained SDS-PAGE gel loaded with samples from each step of hGrx2 purification. The fraction analyzed in each lane is indicated at the top. An identical label for 2 lanes indicates the same fraction from two to three different purifications. 5 μg protein was loaded in each lane. SE, size exclusion; AE, anion-exchange. Arrow indicates molecular weight of Grx2 (16 kD). C, Western blot of hGrx1, hGrx2, and mGrx2 probed with anti-Grx1 primary antibody. hGrx1 shows concentration-dependent immunoreactivity, whereas 4-8 times as much hGrx2 does not react. D, Western blot of hGrx2 with anti-Grx2 antibody, showing concentration-dependent immunoreactivity.

194

195

Figure 3.2. hGrx1 and hGrx2 are selective for protein-GSH mixed disulfide

substrates. Rates of Grx-dependent reduction of [14C] BSA-SS-cysteine ([14C] BSA-

SSC) and [3H] BSA-SS-glutathione ([3H] BSA-SSG) were determined in 0.1M Na/K phosphate buffer, pH 7.5 containing GSH (0.5mM) and 0.1mM RSSG substrate (see

Materials and Methods for details). A, Time-dependent release of [3H] GSSG from [3H]

BSA-SSG. Open circles (○): no hGrx2, closed circles (●) = 9nM hGrx2 (turnover = 39

± 5). B, Time-dependent release of [14C]-cysteine-SSG from [14C] BSA-SSC +/- hGrx2.

Open circles (○): no hGrx2, closed triangles (▲) = 110 nM Grx2 (turnover = 0). Each symbol represents the average of 3-4 experiments ± standard error. When error bars are not visible, they are within the size of the symbol.

196

197

Figure 3.3. Two-substrate kinetic analysis of hGrx2- and mGrx2-catalyzed deglutathionylation demonstrates a ping-pong kinetic pattern. The substrate- dependence of Grx2-mediated reduction of R-SSG was determined using the standard spectrophotometric coupled assay (see Materials and Methods). Reactions were initiated by addition of CSSG (for A-B and E-F), or GSH (for C-D). A-B, Deglutathionylation of

CSSG ± hGrx2 in the presence of fixed GSH with varied CSSG (A); or fixed CSSG with varied GSH (B). Closed diamonds () = 0.25 mM GSH, open diamonds () = 0.3 mM

GSH, closed squares () = 1 mM GSH, open squares () = 2 mM GSH. Closed circles

() = 0.01 mM CSSG, open circles () = 0.02 mM CSSG, closed triangles () = 0.05 mM CSSG, open triangles () = 0.1 mM CSSG. C-D, Deglutathionylation of BSA-SSG

± hGrx2 in the presence of fixed GSH, varied BSA-SSG (C) or fixed BSA-SSG, varied

GSH (D). Closed diamonds () = 0.1 mM GSH, open diamonds () = 0.2 mM GSH, closed squares () = 0.5 mM GSH, open squares () = 1 mM GSH. Closed circles ()

= 5 μM BSA-SSG, open circles () = 10 μM BSA-SSG, closed triangles () = 20 μM

BSA-SSG, open triangles () = 40 μM BSA-SSG. E-F, Deglutathionylation of CSSG ± mGrx2 in the presence of fixed GSH with varied CSSG (E); or fixed CSSG with varied

GSH (F). Closed diamonds () = 0.2 mM GSH, open diamonds () = 0.3 mM GSH, closed squares () = 0.5 mM GSH, open squares () = 0.75 mM GSH. Closed circles

() = 0.02 mM CSSG, open circles () = 0.03 mM CSSG, closed triangles () = 0.05 mM CSSG, open triangles () = 0.1 mM CSSG. Insets, Secondary plots of reciprocal of

Vmax vs. reciprocal of RSSG concentration. Vmax values were calculated from fitting

individual V vs. [S] curves to the Michaelis-Menten equation. Data points represent the

average of at least 3 experiments ± standard error, and when error bars are not visible

198 they are within the size of the symbol. Data in A-B were generated by Amanda Leonberg; results in C-D were generated by David Starke.

199

200

201

Figure 3.4. hGrx2 (C40S) exhibits higher KM and Vmax for CSSG, but lower KM for

GSH, compared to wild type enzyme. Turnover of CSSG by wild type (WT, 135 nM,

closed circles (•)) and C40S (135 nM, closed squares (■)) hGrx2 was measured via the

spectrophotometric coupled assay (see Materials and Methods) with varied CSSG (0.002-

0.5 mM) or GSH (0.02-3 mM), and velocity vs. substrate curves were fit to the

Michaelis-Menten equation. For wild-type hGrx2 enzyme, apparent kcat and KM values for RSSG and GSH are shown in Table 3.2. For hGrx2 (C40S), kcat,app,CSSG = 543 ± 14

-1 -1 min , and KM,app,CSSG = 0.083 ± 0.009 mM; kcat,app,GSH = 589 ± 46 min , and KM,app,GSH =

0.63 ± 0.1 mM. Each point represents the average of at least 3 determinations ± standard

error.

202

203

Figure 3.5. pH rate profile of BSA-SSG deglutathionylation in the absence and

presence of hGrx2. Deglutathionylation of [3H] BSA-SSG was measured at pH 6.5-

10.5, using the standard radiolabel release assay described in Materials and Methods.

Buffers (0.1M) were MES (pH 6.5), HEPES (pH 7-8), HEPPSO (pH 7.5-8.5), and glycine (pH 9-10.5), and the final ionic strength of each buffer was adjusted to 0.3 M using NaCl. Non-enzymatic rates were subtracted from rates in the presence of hGrx2

(15-30 nM), and rates of deglutathionylation ± hGrx2 are expressed as percentages of the maximum rate. hGrx2 concentrations were within the linear range of concentration dependence at each pH. The Figure displays the data from a single experiment that is representative of results from five analogous experiments.

204

Figure 3.6. Determination of the pKa of the active site Cys-SH of hGrx2. hGrx2 (3

μM) was incubated with iodoacetamide (IAM, 0.3 mM) for 3 min in the following incubation buffers: Na citrate, pH 3.5; Na acetate, pH 3.5, 4.0, 4.5, and 5.0; 2-(N-

morpholino)ethanesulfonic acid (MES), 5.0, 5.5, 6.0. All buffers were 10 mM in

concentration, and ionic strength was adjusted to 0.5 M by addition of the appropriate

amount of NaCl or KCl. Following incubation, hGrx2 activity was determined using the

standard spectrophotometric coupled assay (Materials and Methods) after a 50-fold

dilution into the assay mix (0.1 M sodium/potassium phosphate, pH 7.5, 0.2 mM

NADPH, 0.5 mM GSH, 2 U/mL GSSG reductase). Percent activity remaining at each pH

was determined from the ratio of Grx2-mediated deglutathionylation rates for enzyme

preincubated with IAM versus no IAM (see Materials and Methods). Symbols represent

the mean of at least 3 determinations of hGrx1 activity ± IAM preincubation, ± standard error. When error bars are not visible, they are within the size of the symbol. Data generated by Amanda Leonberg.

205

Figure 3.7. Thiyl radical (GS•) scavenging activity of Grx2. A, hGrx (0–1 μM) was

preincubated with 0.5 mM GSH, 0.2 mM NADPH, and 2 U/mL GSSG reductase in 0.1

M Na/K phosphate buffer (pH 7.5) at 30 °C for 5 min (see Materials and Methods and

(33). Then a pre-made complex containing FeCl2 (0.5 mM) and ADP (2.5 mM) was

added to each, and the reactions were initiated by addition of H2O2 (50 μM). Reactions were monitored according to decrease in A340 over 5 minutes (within the linear range of

time-dependence. GSSG formed per min was calculated according to the extinction

coefficient of NADPH, corrected for the plate reader assay (see Materials and Methods).

Inset, region of linear dependence on hGrx concentrations. GS-radical scavenging

activities in this time period were used to calculate turnover (hGrx1 (closed circles, •):

170.0 +/- 7.7; hGrx2 (closed squares, ■): 18.7 +/- 1.2; hGrx1:hGrx2 = 9.1). Symbols

represent the mean of 4-6 determinations ± standard error. B, GSH-dependent peroxidase

activity was measured in 0.1M Na/K phosphate buffer (pH 7.5) containing GSH (0.5

mM) and GSSG reductase (2 U/mL). Reactions were initiated by addition of H2O2 and

A340 was monitored over 5 minutes. Rates of GSSG formation (i.e., NADPH oxidation)

were calculated using the extinction coefficient of NADPH. Glutathione peroxidase

(GPx) was used as a positive control. Closed diamonds (♦): GPx (5nM); open circles (○):

hGrx1 (1 µM); open triangles (Δ): hGrx2 (1µM). Each symbol represents the average of

at least 3 determinations ± standard error. When error bars are not visible, they are

within the size of the symbol.

206

207

CHAPTER 4: GLUTAREDOXIN (GRX1) REGULATES APOPOTOSIS IN CARDIOMYOCYTES VIA NFΚB TARGETS BCL- 2 AND BCL-XL: IMPLICATIONS FOR CARDIAC AGING

Portions of this chapter were included in an article of the same title that is under review for publication at the time of submission of this thesis.

4.1 Abstract

Cardiomyocyte apoptosis is a well-established contributor to irreversible injury following

myocardial infarction (MI), an oxidative insult. Increased cardiomyocyte apoptosis also

is associated with aging in animal models, both at baseline and further in response to MI;

however the mechanism(s) of this increased sensitivity to oxidative stress is not known.

Mixed disulfide formation involving protein cysteine residues and glutathione (i.e.,

protein glutathionylation (protein-SSG)) is known to change the function of signaling

intermediates involved in the regulation of apoptosis. Since glutaredoxin (Grx) is the

thiol-disulfide oxidoreductase enzyme that specifically catalyzes protein

deglutathionylation, we examined its status with aging and its influence on regulation of

apoptosis. We found that Grx1 content and total Grx activity are decreased by 40% in

elderly (24-mo) Fischer 344 rat hearts compared to adult (6-mo) controls. Diminution of

Grx1 to a similar extent in H9c2 cardiomyocytes led to increased apoptotic susceptibility,

decreased NFκB-dependent transcriptional activity, and decreased mRNA and protein

content of the anti-apoptotic NFκB target genes, Bcl-2 and Bcl-xL. Targeted knock- down of Bcl-2 and Bcl-xL in wild-type H9c2 cells to the same extent (i.e., ~50%) as observed in Grx1 knock-down cells increased apoptosis at baseline; and knock-down of

Bcl-xL, but not Bcl-2, also increased oxidant-induced apoptosis to a similar extent as 208 observed in Grx1 knock-down cells. Congruent with the findings for the model Grx1 knock-down cells, natural cardiomyocytes from the Grx1-deficient elderly rats also displayed diminished NFκB-dependent transcriptional activity. Taken together these data support the interpretation that diminished Grx1 in aging animals contributes to increased apoptotic susceptibility via regulation of NFκB and anti-apoptotic target genes.

4.2 Introduction

Cardiomyocyte apoptosis contributes significantly to loss of cardiac function following myocardial infarction (MI), as well as during the development of heart failure, and represents an important therapeutic target in cardiovascular medicine (reviewed in

Mani, 2008; Zidar et al., 2007; Lee and Gustafsson, 2009). Cardiomyocytes from elderly animals exhibit increased apoptotic susceptibility, both at baseline and following oxidative injury (Kajstura et al., 1996; Nitahara et al., 1998; Liu et al., 2002). This increased apoptotic susceptibility may explain the increased risk of heart failure in elderly patients (Kalogeropoulos et al., 2009) and increased post-MI morbidity and mortality in human patients and animal models (Rich et al., 1992; Lesnefsky et al., 1996; Frolkis et al., 1991; Tani et al., 1997; Ashton et al., 2006; Willems et al., 2005). Therefore it is important to characterize molecular mechanisms that contribute to increased apoptosis in aging cardiomyocytes as a prelude to developing therapeutic strategies to minimize cardiac injury in the elderly population. This work examines the role of glutaredoxin in cardiomyocytes as a link between aging and apoptosis.

Human glutaredoxin (Grx1) has been characterized as a thiol-disulfide oxidoreductase enzyme that specifically and efficiently catalyzes protein deglutathionylation (Gravina and Mieyal, 1993). Protein S-glutathionylation is a

209

reversible post-translational modification involving mixed disulfide formation between a

protein cysteine and glutathione (GSH), and it serves both as protection against

irreversible oxidation and as a mechanism of redox signal transduction (reviewed in

Mieyal et al., 2008; Gallogly et al., 2009; Dalle-Donne et al., 2007). Protein

glutathionylation is generally triggered by oxidative stimuli (e.g., growth factor treatment

(Barrett et al., 1999a; Wang et al., 2001b), H2O2 exposure (Chrestensen et al., 2000),

ischemia-reperfusion (Eaton et al., 2002)), but also represents the basal status of certain

protein cysteines in resting cells (e.g., actin (Wang et al., 2001b), mitochondrial Complex

II (Chen et al., 2007), pro-caspase-3 (Pan and Berk, 2007), and IκB kinase (Shelton and

Mieyal, 2008)).

Knowledge is growing about the role of Grx1 as a regulator of apoptosis in

mammalian cells, including cardiomyocytes. The activities of several mediators of

apoptosis have been reported to be modulated by reversible glutathionylation under the

control of Grx1 (e.g., procaspase-3 (Pan and Berk, 2007), p65 (Qanungo et al., 2007),

IKK (Reynaert et al., 2006)). Overexpression of Grx1 in H9c2 cardiomyocytes is cytoprotective, diminishing H2O2-induced apoptosis likely via redox regulation of Akt

(Murata et al., 2003). Likewise increased expression of Grx1 in HEK cells was

protective, increasing survival after glucose deprivation via Grx1 complex formation with

ASK1 (Song et al., 2002). In certain cases, Grx1 has been reported to enhance apoptosis;

e.g., Grx1-catalyzed deglutathionylation (activation) of procaspase-3 enhances apoptosis

in endothelial cells exposed to TNFα and actinomycin D (Pan and Berk, 2007). Thus, the

effect of Grx1 on apoptotic susceptibility seems to be cell type- and stimulus-dependent,

likely reflecting distinct targets of regulation by Grx1 in each case.

210

NFκB is recognized as an important regulator of cell fate in normal and diseased myocardium (reviewed in Jones et al., 2003; Jones et al., 2005), and it has been implicated in the cardioprotective effects of preconditioning (Maulik et al., 1999; Zhao and Kukreja, 2002) and TNF-α stimulation (Mustapha et al., 2000; Bergmann et al.,

2001). Multiple members of the NFκB signaling pathway (e.g., Akt, IKK, p65, and ubiquitin ligase (reviewed in Shelton and Mieyal, 2008) are reported to be inhibited by S- glutathionylation, and thus represent potential targets for regulation by Grx. Indeed, alterations in Grx activity have been linked to changes in NFκB activity in rodent airway

(Reynaert et al., 2006), rat retinal glial (Shelton et al., 2007), and human kidney (Hirota et al., 2000) cell lines.

Here, we report an age-associated diminution of Grx1 in Fischer 344 (F344) rat heart, an observation which prompted mechanistic studies in H9c2 cells to examine the potential contribution(s) of decreased Grx1 to cardiomyocyte apoptosis with aging. We found that diminution of Grx1 in H9c2 cardiomyocytes sensitized them to apoptosis, both at baseline and following oxidative stress, likely via decreased activity of NFκB and subsequent decreased expression of target genes Bcl-2 and Bcl-xL. NFκB-dependent transcription was also decreased in the aging F344 rat heart, suggesting that decreased

Grx1 contributes to age-associated increases in cardiomyocyte apoptosis via an analogous mechanism.

4.2. Materials and Methods

4.2.1. General materials

Unless otherwise specified, all cell culture media, serum, and antibiotics were purchased from Invitrogen , and reagent grade chemicals were obtained from Sigma.

211

4.2.2. Animals

Adult (6-mo) and elderly (24-mo) male Fisher 344 rats were obtained from National

Institute of Aging colonies (Harlan, Indianapolis, IN; Taconic, Germantown, NY) and

housed in the animal facilities of the Louis Stokes Veterans Affairs Cleveland Medical

Center and Case Western Reserve University. Animals were acclimated for at least one week before use, and used for experiments within 4 months of their arrival. The Animal

Care and Use Committees of Case Western Reserve University and the Louis Stokes

Veterans Affairs Cleveland Medical Center approved all animal protocols.

4.2.3. Specific Methods

Culture and maintenance of H9c2 cells – H9c2 cells (rat embryonic cardiomyocytes) (a gift from Laura Nagy, Cleveland Clinic Foundation) were cultured in DMEM (Invitrogen

#11965-092) with 10% certified fetal bovine serum (FBS) and 1X

“antibiotic/antimycotic” (1U/mL penicillin, 1 μg/mL streptomycin, and 0.025 ng/mL amphotericin-B) at 37°C, 5% CO2. Cells were routinely passaged after reaching

confluence. All experiments utilized cells of passage number 10-30.

Knock-down of Grx1, Bcl-2, and Bcl-xL in H9c2 cells –2-2.5 x 104 H9c2 cells were seeded onto each well of 6-well culture dishes containing 2mL DMEM (containing 10%

FBS and 1X antibiotic/antimycotic). The following day, cells were rinsed twice with serum-free medium and transfected with control (non-targeting) or Grx1-, Bcl-2-, or Bcl- xL-targeted siRNA (25-100nM, Thermo Scientific) in the presence of oligofectamine (3

µL/well, Invitrogen) according to the manufacturer’s instructions. Four hours later, 50

µL DMEM containing 30% serum (no antibiotics) was added to each well. Cells were analyzed for protein content and/or susceptibility to apoptosis 24-48 h following the end

212

of transfection. For each siRNA target, the extent of knock-down was adjusted by

varying the dose of siRNA and determining the percentage of knock-down by Western

blot analysis (see below) over a 3-day period post-transfection.

Stable knock-down of Grx1 in H9c2 cells – Each of the following DNA sequences were

subcloned individually into the p.SUPER.retro.puro vector (Oligoengine) using T4 DNA ligase: (1) 19 random residues corresponding to no known gene in the rat or mouse genome; (2) base pairs 94-112 of the rat Grx1 gene; (3) rat Grx1 base pairs 231-249; or

(4) rat Grx1 base pairs 300-318. For transfection, 6 μg of Grx1-coding plasmid DNA (2

μg of each construct, for knock-down cells) or 2 μg of plasmid containing the scrambled

sequence (for control cells) was used to transfect 6 million BOSC cells (kindly provided

by Dr. George Stark, Cleveland Clinic Foundation) plated on a 100mm dish in serum-free

DMEM using Lipofectamine and PLUS reagents (Invitrogen), according to the

manufacturer’s instructions. After incubation for 4 h at 37 C, 5% CO2, medium was removed by aspiration and replaced with DMEM containing⁰ 10% heat-inactivated FBS and 1X antibiotic/antimycotic. After 24 h (day 2), medium was collected from BOSC cells and diluted 1:1 with DMEM (10% FBS, 1X antibiotic/antimycotic). After addition of polybrene (5-dimethyl-1,5-diazaundecamethylene polymethobromide, hexadimethrine

bromide) (10 μg/mL final concentration), the mixture was sterile-filtered and added to

100,000 H9c2 cells plated on a 100 mM dish. Fresh medium was replaced on the BOSC

cells, and this procedure was repeated on days 3 and 4. On day 5, 2 μg/mL puromycin

(Sigma) was added to the H9c2 cell medium. Cells were selected for approximately 7

days and analyzed for Grx1 content and total Grx activity. Grx knock-down was

213

maintained for weeks in the presence of puromycin and H9c2 cells up to 6 passages post-

transfection were used for experiments.

Determination of Grx activity – For measurement of Grx activity in F344 rat heart tissue,

rat myocardium (250 mg) from adult (6-10-mo) or elderly (24-28-mo) F344 rats was homogenized at 4 C in 1 mL buffer containing potassium phosphate (10 mM), pH 7.5;

EDTA (5 mM); phenylmethane⁰ -sulfonyl fluoride (50 μM); and 2 mM β-mercaptoethanol using a glass homogenizer. The homogenate was centrifuged at 4 C sequentially at 700,

2000, and 10,000 xg for 10 min at each rate. Supernatants were dialyzed⁰ overnight at 4

C against the same buffer at a ratio of 1mL supernatant:1000 mL dialysis buffer (with 2 changes⁰ of buffer) to remove GSH. Grx activity was measured by the radiolabel assay, as described by Chrestensen et al. (2000). Briefly, 10-50 µL cell lysate was preincubated in sodium/potassium phosphate buffer (0.1 M, pH 7.5) containing GSH (0.5 mM) for 5 min in a 30 C water bath. Reactions were initiated by addition of the substrate [35S]

BSA-SSG (0.1⁰ M final concentration (synthesis described in Gallogly et al., 2008;

Chapter 3). Aliquots of the reaction mixture were removed at t = 10 s, 1 min, 2 min. and

3 min and quenched by addition of an equal volume of ice cold TCA (20%). Precipitated

protein was pelleted by centrifugation for 5 min at 10,000 xg at 4 C, and supernatants

were analyzed for released radiolabel via scintillation counting.⁰ Grx-mediated

deglutathionylation rates were calculated by subtracting turnover in the absence of Grx

from turnover in the presence of enzyme, and using the specific radioactivity of the

released glutathionyl moiety (determined separately) to calculate nmol product per min

per mg protein. TR and GR activities were determined as described by Starke et al.

(1997).

214

For measurement of Grx activity in cardiomyocytes isolated from F344 rats

(described below), cell pellets were homogenized in NP-40 lysis buffer (50 mM Tris-

HCl, pH 8, 150 mM NaCl, 1% NP-40) in disposable, hand-held homogenizers (Fisher) and cleared by centrifugation at 10,000 x g for 5 min at 4°C, then activity was measured spectrophotometrically. Approximately 60 µL of the supernatant was added to microwells of a 96-well plate containing a mixture of 0.1 M Na/K phosphate buffer, pH

7.5, 0.5 mM GSH, 0.2 mM NADPH, and 2 U/mL GSSG reductase (180 µL, total volume), and incubated for 5 min at 30°C. Reactions were initiated by addition of 20 µL of the substrate cysteine-SSG (0.1 mM final, Toronto Research Chemicals) and rates of deglutathionylation were monitored by change in A340nm/min as described by Srinivasan et al. (1997). Activity was normalized to protein content measured by the BCA assay

(Pierce).

Grx activity in H9c2 cells was measured spectrophotometrically, following trypsinization of control and Grx1 knock-down cells, lysis in NP-40 lysis buffer on ice for 10 min, and clearing of lysates by centrifugation at 10,000 g (4°C). Supernatants (5-

20 μL) were analyzed for Grx activity and for protein content as described above. For all analyses, Grx activity was determined within the linear range of concentration dependence of activity on volume (μL) of lysate added.

Western blotting – H9c2 cells were collected as described for determination of Grx activity, except that Sigma Protease Inhibitor Cocktail (5%, final) was included in the lysis buffer. The cytosolic fraction of F344 rat heart homogenate was prepared as described above for analysis of Grx activity. Lysates/homogenates containing 10-100 µg of protein were incubated with 4X sample buffer (100 mL 0.5 M Tris-HCl, pH 6.8, 80

215

mL glycerol, 160 mL 10% SDS, 20 mL 1% bromophenol blue; final concentrations at

1X: 34 mM Tris-HCl, pH 6.8, 5.5% glycerol, 1.1% SDS, 0.014% bromophenol blue) and

DTT (10 mM, final) for 10 min. at 95°C; then, IAM (25 mM, final) was added and

samples were heated at 95°C again for 5 min. Samples were resolved on 12.5%

polyacrylamide gels and transferred to PVDF membranes. Membranes were blocked

with nonfat milk (5% in Tris-buffered saline, TBS) for 1 h at room temperature, then with

primary antibody in 5% milk/TBS overnight at 4°C. Membranes were washed with TBS

(3 x 10 min), then incubated with the appropriate secondary antibody (1:10,000) in 5%

milk/TBS for 1 h and washed again. ECL or Pico Supersignal developing reagent

(Pierce) was used for chemiluminescent detection of antibody binding. Primary

antibodies were anti-Grx1 (Steven A. Gravina, PhD thesis, Case Western Reserve

University, 1993), 1:1000), anti-actin (Sigma, clone AC-74, 1:60,000), anti-GAPDH

(Calbiochem, 1:100,000), anti-Bcl-2 (BD Biosciences, 1:500), anti-Bcl-xL (Cell

Signaling Technologies, 1:5000), anti-p50 (Santa Cruz, sc-114, 1:1000), anti-p65 (Santa

Cruz, sc-372, 1:2000). Secondary antibodies were anti-rabbit or anti-mouse (both from

Jackson Laboratories). For detection of Bcl-2 and Bcl-xL, 0.1% Tween (final) was added to the TBS to minimize background signal.

Determination of Trx1 content – Heart apices from F344 rats were quick-frozen in liquid

N2, then thawed and homogenized for 20 s in homogenization buffer (20 mM Tris, pH

7.5, 1% Triton X-100, 100 mM NaCl, 40 mM NaF, 1 mM EDTA, 1 mM EGTA, 5%

Sigma Protease Inhibitor Cocktail) using a Polytron homogenizer (Brinkmann) set at

70% maximum speed. Homogenates were incubated on ice for 30 min, then centrifuged

at 10,000 x g for 10 min. Protein content of supernatants was determined by the BCA

216

assay (Pierce) and Trx1 content was analyzed by Western blot analysis as described

above. (The polyclonal sheep, anti-human Trx1 antibody (QCB) was generated from

peptide sequences of the Trx1 protein, and used at a dilution of 5 μg/mL). TR and GR

activities were measured on heart tissue homogenates as described by Starke et al.

(1997).

Real-time PCR – Cells transfected with control or Grx1 siRNA for 48h were collected in

a total volume of 1mL Trizol (Invitrogen) per 6 wells of a 6-well culture dish, and RNA

was isolated from each sample using the RNeasy kit (Qiagen). Relative quantities of

mRNA were then determined in two different ways. In the first, cDNA was generated

from each sample according to the ABI High Capacity RT kit protocol (200 ng RNA

input), and real-time PCR was performed on an Applied Biosystems 7500 Fast Real Time

PCR system according to the manufacturer’s protocol, using primers for 18S rRNA or rat

Bcl-2 (Applied Biosystems). To confirm the diminution of Bcl-2 mRNA in Grx1 knock- down cells, and to determine the effect of Grx1 knock-down on content of Bcl-xL mRNA, relative quantities of Bcl-2 and Bcl-xL mRNA were determined simultaneously using a PCR-array from SA Biosciences. cDNA from control and Grx1 knock-down cells was generated from Trizol lysates using the SA Biosciences RT2 First Strand Kit

(500 ng input), and samples were loaded onto a 384-well plate containing probes for 84

apoptosis-related genes and 6 housekeeping genes (Cat. No. PARN-012, SA

Biosciences). PCR analysis was performed on an ABI 7900 Real-Time PCR system

(Applied Biosystems) using SYBR Green/ROX Master Mix (Cat. No. PA-012-8, SA

Biosciences). Cycle conditions were: (1) hold at 95°C for 10 min; (2) 40 cycles of 15 s at 95°C followed by 1 min at 60°C. Relative changes in gene expression were calculated

217

using the ΔΔCt method (Livak and Schmittgen, 2001). Western blot analysis indicated

that protein content of GAPDH did not change with Grx1 knock-down; thus, it was used

to normalize the calculated relative quantities of mRNA. Duplicates of each sample were

averaged prior to normalization and calculation of relative expression. A total of 3 pairs

of samples were analyzed, and Grx1 knock-down was confirmed separately for each set

of samples by Western blot analysis.

H2O2 treatment of H9c2 cells – H9c2 cells were incubated in culture medium with or

without H2O2 (Fisher Scientific, 400μM final) for 5 min at 30 °C/5% CO2. Following

treatment, the medium on all cells was replaced with fresh medium without H2O2. For wild-type H9c2 cells, treatments were performed in medium containing 10% serum and

1X antibiotic/antimycotic (normal culture medium). For H9c2 cells transfected with siRNA, treatment was done in medium containing 1.4% serum (the final serum concentration following transfection) without antibiotic/antimycotic. H2O2 dose-response experiments on wild-type cells showed no difference in apoptosis in wild-type cells

treated with H2O2 in 1.4%- vs. 10%-serum medium (data not shown). 24 h after treatment, cells were analyzed for apoptosis by Hoechst staining (see below).

Simulated ischemia/reperfusion of H9c2 cells –H9c2 cells were incubated with glucose-

free Krebs-Ringer-Henseleit (KRH) buffer (155 mM NaCl, 5 mM KCl, 1mM CaCl2, 1

mM KH2PO4, 1.2 mM MgSO4, 25 mM HEPES, pH 6.2 or pH 7.4 (Kim et al., 2006)).

Cells at pH 6.2 were placed in a plexiglass chamber (Billups-Rottenberg, Del Mar, CA)

which was made hypoxic by sparging with 95% N2, 5% CO2, 0.5% O2 for 4 min at 2.5

psi. The chamber was placed in the same incubator as the one containing the cells at pH

7.4 (normoxic condition), and all cells were incubated for 3 h. Following simulated

218 ischemia, hypoxic H9c2 cells were removed from the chamber and the medium on all cells was replaced with KRH buffer, pH 7.4. Cells were incubated for 2 h more under normoxic conditions to simulate reperfusion, then analyzed for apoptosis by Hoechst staining (see below).

Determination of apoptosis – H9c2 cells were incubated with Hoescht 33342 dye

(Invitrogen, H1399, 1 µg/mL final) for 10 min and nuclear condensation and fragmentation were visualized with a Leica fluorescence microscope equipped with a

DAPI filter. The percentage of apoptotic cells was calculated by dividing the number of cells with condensed and/or fragmented nuclei by the total number of cells counted in each well of a 6-well plate. At least 300 cells were counted in each well, and each well represented one determination of a triplicate experiment for each treatment. All cell counting was performed in a blinded fashion.

NFκB activity in H9c2 cells – 24 h after transfection with siRNA (control or Grx1), H9c2 cells were co-transfected for 12 h with 1µg of 5X-NFκB Luciferase plasmid (Stratagene) and 0.1 µg of Renilla plasmid (Promega) according to the Lipofectamine (Invitrogen) reagent protocol. 24 h after transfection, Luciferase production was assayed using the

Dual-Luciferase assay system (Promega) and measuring emission with a luminometer

(Molecular Devices fitted with SOFTmax PRO software). NFB activity is expressed as the ratio of firefly luciferase-dependent luminescence to Renilla luciferase-dependent luminescence.

Effect of NFκB inhibition on apoptotic susceptibility –25,000 cells were seeded on each well of a 6-well dish in regular medium without antibiotics, then cultured for 24h. Cells were then co-transfected with 1 μg/well NFκB-Luciferase plasmid (Stratagene) and 0.1

219

μg/well Renilla plasmid (Promega) using the Lipofectamine 2000 transfection reagent

(Invitrogen) using a ratio of 1μg DNA:2μL Lipofectamine 2000. Approximately 8 hr

after the start of transfection, medium was replaced with antibiotic-free culture medium

containing BMS 345541 (4(2-aminoethyl)amino-1,8-dimethylimidazo(1,2-a)quinoxaline)

(0-20 μM) and cells were cultured overnight. The next morning (24h after transfection,

~14h after BMS treatment), cells were either collected for determination of NFκB

activity (Luciferase assay, see above) or devoted to determination of apoptotic

susceptibility. Of the latter cells, half were treated with H2O2 (400 μM x 5 min, followed

by replacement of H2O2-containing media with regular culture medium) and half were

subjected only to a change of medium (no H2O2 treatment). 24h after H2O2 treatment,

apoptosis was assessed by Hoechst staining as described above. Note: For BMS-treated

cells, Firefly-luciferase activity was normalized to protein content (measured by

absorbance at 280 nm) rather than Renilla-luciferase activity because BMS treatment

appeared to interfere with measurement of Renilla activity.

Isolation of primary cardiomyocytes – Cardiomyocytes from adult and elderly F344 rats

were isolated as described by Patel et al. (2006), with modifications designed to

maximize the yield of elderly myocytes. First, animals were heparinized (3,000 units ip)

5 min prior to administration of pentobarbital (100 mg/kg). Hearts were removed, placed

in ice-cold heart media (112 mM NaCl, 5.4 mM KCl, 1 mM MgCl2, 9 mM NaH2PO4,

11.1 mM D-glucose, 10 mM HEPES, 30 mM taurine, 2 mM DL-carnitine, 2 mM creatine) to remove blood; then transferred to cardioplegic solution (heart media containing 20 mM KCl) for cannulation. Hearts were cannulated via the aorta and perfused in a retrograde fashion on a Langendorff apparatus (flow rate ~ 6 mL/min) with

220

heart media for 5 min, followed by heart media containing Collagenase Type II

(Worthington Biochemical, ~400 U/mL; lot chosen for its effectiveness with F344 rats) for 20 min. Following digestion, the ventricles were removed and minced for 3-4 min in

10 mL collagenase-free heart media to minimize damage to the fragile elderly myocytes.

Following mincing, cardiomyocytes were triturated for 3 min with a disposable plastic pipettor and filtered through an 80 µm steel mesh screen. The filtrate volume was brought to 25 mL by addition of ~15 mL wash buffer (heart media containing 1% BSA and 20 µM CaCl2), then centrifuged at low speed (~170 x g) for 25 s. 20 mL of the supernatant was then removed, 25mL wash buffer was added to the loose pellet, and the tube was gently inverted 2-4 times. After 10 min settling at room temperature, the

centrifugation and resuspension was repeated for a total of 2 washes. Ca2+ was

reintroduced to cells in wash buffer by 2 stepwise additions (decreased from four

additions to minimize myocyte manipulation) of 150 μL CaCl2 solution (100 mM stock)

to achieve a final Ca2+ concentration of 1.0 mM (reported to be better tolerated by cardiomyocytes from elderly rats (Lieber et al., 2004) than the typical concentration of

1.2 mM used for isolation of cardiomyocytes from neonatal or adult animals). After each addition of CaCl2, cells were gently mixed and allowed to settle for ~10 min. After the

second settling, the cell supernatant (wash buffer containing 1.0 mM Ca2+) was replaced

with plating medium (Medium 199 with Hank’s salts, 4% FBS, 1%

penicillin/streptomycin). Yields were typically ~1-2 x 106 cells (~50% rods) from adult

hearts, and 2.5-5 x 105 cells (10-30% rods) from elderly hearts. Comparison of Grx activity in adult and elderly myocytes indicated that the myocyte preparations exhibit the same age-associated decline in Grx activity observed in heart tissue (see Results),

221

suggesting that representative populations of cardiomyocytes were isolated from rats of

each age group.

Determination of NFκB activity in primary cardiomyocytes – Following isolation,

cardiomyocytes in plating medium (Medium 199 containing Hank’s salts, FBS (4%), and

penicillin/streptomycin (1%)) were allowed to settle by gravity, then resuspended and

divided into 3-6 (for elderly rats) or 12-24 (for adult rats) equivalent volumes, according

to the estimated yield (i.e., amount of settled cells). Each sample was pelleted by brief

centrifugation at ~175 x g for 10-20 s, supernatants (containing medium and fibroblasts)

were removed by aspiration, and each pellet was resuspended in 200 µL maintenance

medium (4 parts Medium 199 with Hank’s salts, BSA (1%) and penicillin-streptomycin

(1%); 1 part Joklik Modified Medium (Sigma)) containing Renilla (MOI = ~20) and

NFκB-Luciferase (MOI = ~100) or MCS (negative control, MOI = ~100) adenovirus

(Vector Biolabs) and incubated at 30°C/5% CO2 for 1 h. Following incubation, each

mixture of cardiomyocytes in virus-containing medium was diluted into 2mL

maintenance medium and added to one well of a 6-well tissue culture plate pre-coated with mouse laminin (Invitrogen). For elderly rats, cardiomyocytes from 2 hearts were plated sequentially onto the same 3-6 wells to achieve comparable cell density and to ensure sufficient yield for the Luciferase assay. Cardiomyocytes were cultured for 24h, then medium was removed and 150 µL 1X Passive Lysis Buffer (Promega) was added to each well. Plates were rocked for 15 min at room temperature; then the cells were scraped, quick-frozen in Eppendorf tubes in a dry ice/EtOH bath, and stored at -80°C. 20

µL of each sample was used to determine Luciferase and Renilla activities using the

Dual-Luciferase assay system (Promega) as described above. For each preparation of

222

myocytes, at least 3 replicate determinations were performed and then averaged to

represent a single experiment (i.e., n number). Cells infected with MCS adenovirus

exhibited no Luciferase activity above background (i.e., Luciferase and Stop & Glow

reagents alone, data not shown).

Statistical analysis – Unless otherwise indicated, data are reported as mean ± standard

error (mean ± SEM). Individual data points more than two standard deviations from the

mean were considered to be outliers and omitted. Differences between data sets were

tested for statistical significance using Student’s t-test (Microsoft Excel) on primary data.

4.3. Results

Grx1 content and total Grx activity are decreased with aging in the F344 rat heart:

Cytosolic fractions of heart tissue from adult (6-10 mo.) and elderly (24-28 mo.) F344

rats were analyzed by semi-quantitative Western blotting and revealed an age-associated

decrease in Grx1 content of approximately 40% (Figure 4.1A, left, p. 236). Total Grx

activity (i.e., GSH-dependent deglutathionylation of protein-SSG substrate) was

diminished by a similar margin with aging (Figure 4.1A, right, p. 236). In contrast,

thioredoxin 1 (Trx1) content and thioredoxin reductase (TR) and glutathione disulfide

reductase (GR) activities were unchanged with aging; i.e., Trx1 content was

indistinguishable by western blot analysis compared to standards, and thioredoxin

reductase (TR1) activity was unchanged (6-mo: 1.68 ± 0.13 mol/min/ml (n = 4); 24 - mo: 1.63 ± 0.08 (n = 4)). GR activity for cytosolic samples from 6-mo and 24-mo rats was 14.5 ± 0.7 nmol/min/mg (n = 2) and 16.7 ± 0.04 nmol/min/mg (n = 2).

To confirm that the age-associated diminution of Grx1 occurs in cardiomyocytes

(which contribute the majority of cardiac protein to heart tissue), cardiac myocytes were

223

isolated from adult and elderly F344 rats via adaptation of the method of Patel et al.

(2006) (see Materials and Methods). Although preparations from elderly animals yielded

fewer cells than those from adult animals (0.3-0.5 x 106 cells for elderly vs. 1-2 x 106

cells from adults), both preparations, after plating on laminin and changing the medium,

yielded mostly adherent rod-shaped cardiomyocytes with similar overall morphology

(Figure 4.1B-C, p. 236). Analysis of lysates from isolated cardiomyocytes indicated a

similar diminution in total Grx activity as observed in tissue cytosol (Figure 4.1D, p.

236).

Knock-down of Grx1 in H9c2 cells: To investigate the specific contribution(s) of

decreased Grx1 to increased apoptotic susceptibility in cardiomyocytes, Grx1 was

knocked-down using siRNA in H9c2 cells (rat embryonic cardiomyocytes). siRNA dose

and post-incubation time were adjusted to achieve a diminution of Grx1 in H9c2 cells

that was similar to the decrease observed in aging cardiomyocytes (Figure 4.2A, p. 238).

Thus, in H9c2 cells transfected with Grx1-targeted siRNA, Grx1 content and activity

(Figure 4.2B, p. 238) were decreased by approximately 50% compared to wild-type and to control cells (transfected with a pool of non-targeted siRNA). A comparable diminution of Grx1 content and activity was achieved when Grx1 was knocked-down stably using shRNA, i.e., Grx1 content in shRNA knock-down cells was 31 ± 16% lower than in cells infected with scrambled (control) shRNA; total Grx activity was diminished by 40 ± 5% in shRNA knock-down cells compared to control cells (Data generated by

Dr. Harish Pai).

Grx1 knock-down increases oxidant-induced apoptosis in H9c2 cells: To test the potential contribution of diminished Grx1 on susceptibility to apoptosis following an

224

oxidative insult, control and Grx1–deficient H9c2 cells were subjected to simulated

ischemia reperfusion (sIR) (Kim et al., 2006), or treated with H2O2, previously shown to

trigger a classical apoptotic cascade in H9c2 cells (Han et al., 2004). Following treatment, apoptosis was assessed by Hoechst staining. Grx1-deficient cells exhibited increased apoptosis compared to control cells, both at baseline and following oxidative stress; i.e., the percentage of apoptotic cells was approximately doubled in Grx1-deficient cells compared to control cells following simulated IR or H2O2 treatment (Figure 4.3A, p. 240). Analogous results were observed in H9c2 cells with stable knock-down of Grx1

(Figure 4.3B, p. 240).

NFκB transcriptional activity is decreased in Grx1-deficient cells: We hypothesized that decreased activity of the NFκB pathway contributes to the increased apoptotic susceptibility of Grx1-deficient cells. To determine whether Grx1 knock-down results in decreased transcriptional activity of NFκB, NFκB-dependent transcription was measured

in control and Grx1-deficient cells using a standard dual-Luciferase reporter assay.

Indeed, NFκB transcriptional activity was partially diminished (37 ± 11% decrease) in

Grx1 knock-down cells compared to control cells (Figure 4.4C, p. 242). This decrease is

unlikely to reflect decreased content of NFκB transcription factor subunits, since Western

blot analysis indicated no statistically significant difference in the amount of total p50

and p65 in control vs. Grx1 knock-down cells (Figure 4.4A-B, p. 242).

Inhibition of NFκB-dependent transcription is sufficient to sensitize H9c2 cells to

oxidant-induced apoptosis: To determine whether diminution of NFκB activity by a

similar extent as seen in Grx1-deficient cells is sufficient to increase apoptotic

susceptibility in H9c2 cells, wild-type H9c2 cells were treated with the IKK inhibitor

225

BMS 345541 (Burke et al., 2003). Both NFκB activity and apoptotic susceptibility were

assessed following inhibitor treatment. BMS treatment resulted in dose-dependent

diminution in NFκB activity (Figure 4.5A, p. 244), as well as increased susceptibility to

apoptosis compared to untreated cells, in the presence and absence of H2O2 (Figure 4.5B,

p. 244). Importantly, the concentration of BMS at which NFκB-dependent transcription

was decreased by ~30% (i.e., the amount of diminution observed in Grx1-deficient cells) was sufficient to increase apoptotic susceptibility, suggesting that ~30% inhibition of the

NFκB pathway could contribute to the increased apoptotic susceptibility of Grx1- deficient cells. mRNA and protein content of Bcl-2 and Bcl-xL are decreased in Grx1-deficient cells:

Anti-apoptotic NFκB target genes Bcl-2 and Bcl-xL promote cellular survival in many cell types, including cardiomyocytes (reviewed in Gustafsson and Gottlieb, 2007; Gross et al., 1999; Kim, 2005). In transgenic mouse models, Bcl-2 and Bcl-xL protect cardiomyocytes from apoptosis and improve cardiac function, particularly in the contexts of ischemia-reperfusion injury and heart failure (Chen et al., 2002; Huang et al., 2003;

Chen et al., 2001; Brocheriou et al., 2000; Imahashi et al., 2004; Weisleder et al., 2004;

Moorjani et al., 2007). Conversely, decreases in Bcl-2 and/or Bcl-xL are associated with increased cardiomyocyte apoptosis in experimental models of ischemia (Lazou et al.,

2006; Tantini et al., 2006; Bonavita et al., 2003). We hypothesized that the increased apoptotic susceptibility in Grx1-deficient cells was due to decreased NFκB-dependent transcription of Bcl-2 and/or Bcl-xL. Indeed, real-time PCR analysis indicated that relative quantities of Bcl-2 and Bcl-xL mRNA were substantially decreased in Grx1- deficient cells (Figure 4.6, p. 245). Western blot analysis confirmed that contents of the

226

Bcl-2 and Bcl-xL proteins were also diminished substantially (~50%) with knock-down

of Grx1 (Figure 4.7, p. 247).

Effect of decreased Bcl-2, Bcl-xL, or both on oxidant-induced apoptosis in H9c2 cells:

To determine the relative contributions of decreased Bcl-2 and Bcl-xL in H9c2 cells, the proteins were knocked down individually or together (Figure 4.8A-D, p. 249), and apoptotic susceptibility was assessed both at baseline and following H2O2 treatment

(Figure 4.8E, p. 250). Knock-down of Bcl-2 or Bcl-xL in H9c2 cells increased apoptosis

at baseline, each to a level comparable with that observed in Grx1 knock-down cells. In

contrast, only knock-down of Bcl-xL increased oxidant-induced apoptosis in H9c2 cells, with the percentage of apoptotic cells closely matching that observed with Grx1 knock- down. Importantly, knock-down of Bcl-2 or Bcl-xL did not affect expression of the other protein (Figure 4.8C, p. 249). Simultaneous knock-down of Bcl-2 and Bcl-xL resulted in an apoptotic phenotype that was statistically indistinguishable from Bcl-xL knock- down alone.

NFκB activity in cardiomyocytes from adult and elderly F344 rats: Cytosolic extracts prepared from elderly (24-28-mo) F344 rat heart tissue exhibited decreased Grx1 content and total Grx activity compared to adult control samples (6-10-mo), and a similar diminution in Grx activity was observed in lysates of cardiomyocytes isolated from elderly vs. adult F344 rat hearts (see Figure 4.1, p. 235). To determine whether NFκB activity is decreased also in elderly cardiomyocytes (as with Grx1 knock-down in H9c2 cells), NFκB-dependent transcription was measured in cardiomyocytes isolated from adult and elderly F344 rats. Indeed, cardiomyocytes from elderly F344 rats exhibited

227

substantially lower NFκB-dependent transcriptional activity compared to adult controls

(activity = 30 ± 9% of the adult control value, Figure 4.9, p. 251).

4.4 Discussion

The current study examined the role of glutaredoxin (Grx1) in modulation of

apoptosis in cardiomyocytes, and identified a NFB -dependent mechanism by which

diminution of Grx1 may contribute to increased apoptotic susceptibility with aging. We

found that Grx1 content and total Grx activity are decreased with aging in heart tissue

and in cardiomyocytes isolated from Fischer 344 rats, a well-established animal model of aging, whereas other thiol-disulfide oxidoreductase enzymes were found to be unchanged. Based on observations that many apoptotic mediators are subject to regulation by reversible glutathionylation (Pan and Berk, 2007; Qanungo et al., 2007;

Reynaert et al., 2006), and that overexpression of Grx1 protects H9c2 cardiomyocytes from oxidant-induced apoptosis (Murata et al., 2003), we hypothesized that the age- related decrease in Grx1 contributes to the increased apoptotic susceptibility already documented in cardiomyocytes from elderly animals (Kajstura et al., 1996; Nitahara et al., 1998; Liu et al., 2002). Therefore we knocked down Grx1 in H9c2 cells to a similar extent as in cardiomyocytes from elderly rats to assess the independent contribution of diminished Grx1 to apoptotic susceptibility.

Selective diminution of Grx1 indeed led to an increase in apoptotic cells at baseline as well as following oxidative insults previously shown to trigger cardiomyocyte apoptosis

(Han et al., 2004; Kim et al., 2006). It is noteworthy that both transient (siRNA) or stable (shRNA) knock-down of Grx1 led to analogous increases in oxidant-induced apoptotic susceptibility of the model cardiomyocytes, because the latter approach is more

228

akin to the decrease in Grx1 in the cells from elderly animals. Transcriptional activity of

NFκB, a transcription factor with anti-apoptotic effects in cardiomyocytes (Mustapha et al., 2000; Bergmann et al., 2001), was concomitantly decreased in both Grx1 knock- down cells and in primary cardiomyocytes isolated from elderly F344 rats.

To determine whether decreased NFκB activity could account for the apoptotic

phenotype of Grx1-deficient cells, the effect of NFκB inhibition on apoptotic

susceptibility was tested in wild-type H9c2 cells. Thus, treatment of H9c2 cells with the

IKK inhibitor BMS 345541 (Burke et al., 2003) led to a dose-dependent diminution of

NFκB-transcriptional activity, which correlated with increased apoptosis both at baseline

and following H2O2 treatment. Importantly, diminution of NFκB transcriptional activity

to a similar extent as observed in Grx1 knock-down cells was sufficient to increase

apoptotic susceptibility, implicating the decreased NFκB activity in the apoptotic

phenotype of Grx1 knock-down cells.

mRNA and protein contents of the anti-apoptotic NFκB transcription products Bcl-2

and Bcl-xL were decreased by about 50% in Grx-deficient cells. Knock-down of Bcl-2

and/or Bcl-xL in H9c2 cells to a similar extent (i.e., ~40-60%) increased apoptosis at

baseline, suggesting that diminished Bcl-2 and Bcl-xL contribute to the increased

apoptotic susceptibility of resting Grx1-deficient cells. In remarkable contrast, knock-

down of Bcl-2 alone had no effect on oxidant-induced apoptosis in H9c2 cells, whereas

diminution of Bcl-xL alone (or in combination with Bcl-2) increased oxidant-induced

apoptosis to the same extent as observed in Grx1 knock-down cells, fully accounting for

the apoptotic phenotype of these cells. Taken together, these observations suggest a

mechanism for the increased apoptotic susceptibility of aging cardiomyocytes in which

229

diminished Grx1 leads to decreased NFκB activity, resulting in decreased content of the

anti-apoptotic proteins Bcl-2 and Bcl-xL, and a lower threshold for commitment to

apoptosis.

Control of apoptotic susceptibility by Grx1 via regulation of NFκB: , Grx1 has been

implicated in the regulation of apoptosis via modulation of the glutathionylation status of

apoptotic mediators, including NFκB (Shelton and Mieyal, 2008). S-glutathionylation is

inhibitory towards many NFκB signaling intermediates, (e.g., IKK (Reynaert et al.,

2006), p50 (Pineda-Molina et al., 2001), p65 (Qanungo et al., 2007), 20S proteasome

(Demasi et al., 2003), ubiquitin-activating (Jahngen-Hodge et al., 1997) and -carrier

proteins (Obin et al., 1998)), and increased expression of Grx1, the primary intracellular deglutathionylating enzyme, is correlated with increased NFκB transcriptional activity

(Hirota et al., 2000; Reynaert et al., 2006; Shelton et al., 2007). Thus, in most cases,

Grx1 activity is expected to support NFκB activity, likely via deglutathionylation of one

or more mediators in the NFκB signaling pathway. A notable exception is in hypoxic

pancreatic cancer cells, where Grx1 enhances the glutathionylation of p65, leading to

decreased DNA binding and diminished NFκB transcriptional activity (Qanungo et al.,

2007).

We observed decreased NFκB activity in Grx1 knock-down H9c2 cells as well as in

Grx1-deficient (elderly) primary cardiomyocytes. Thus, the likely mechanism of NFκB

regulation by Grx1 in these cardiomyocytes is direct deglutathionylation (activation) of

one or more components of the NFκB axis, so that diminution of Grx1 activity leads to

increased glutathionylation of one or more control points in the pathway and

deactivation. The specific site(s) of regulation of the NFκB pathway in the Grx1 knock-

230

down H9c2 cells (and in the aging heart) is not known, and identifying these sites

represents a natural direction for future studies. Besides the signaling intermediates

described above, other possibilities include upstream activators (e.g., Akt (Murata et al.,

2003), Ras (Adachi et al., 2004a)) or inhibitors (e.g., PTEN (Cruz et al., 2007)). Other

investigators have presented evidence for activation of Akt activity in H9c2 cells and

heart tissue by Grx1 (Murata et al., 2003; Malik et al., 2008), but the relationship

between those studies that involved overexpression of Grx1 and the effects of Grx1

diminution in aging remains to be elucidated.

To our knowledge, this is the first report of diminished NFκB activity in the aging

heart, documented in cardiomyocytes from adult and elderly rats. Age-associated changes in NFκB activity have been reported in other contexts. For example, enhancement of NFκB-dependent transcription was detected with aging in rat arteries,

human liver and spleen, and multiple mouse tissues (Ungvari et al., 2007; Adler et al.,

2007). A recent report (Gao et al., 2008) concluded that NFκB activity was increased

with aging in cardiac tissue from Sprague-Dawley rats; however, it appears that this

conclusion was based on increased content of p65 in the nucleus, which is necessary but

not sufficient for NFκB transcriptional activity. Age-associated decreases in NFκB activity have been reported in mouse liver, as well as in neurons and gingival tissue from elderly rats (Okaya et al., 2005; Patel and Brewer, 2008; Kim et al., 2009), while no age-

related change in NFκB transcriptional activity was reported in a recent study of rat

muscle precursor cells (Lees et al., 2009). To date, studies of the effects of aging on

NFκB activity appear to reflect tissue- and organism-specific differences in regulation.

231

Studies of thiol-disulfide oxidoreductase systems with aging yield a complex picture,

possibly reflecting tissue-, organism- and sex-specific mechanisms of regulation. We

found that Grx1 content and total Grx activity were decreased by about 40% in cytosolic

extracts from elderly F344 rat hearts, while Trx, TR, and GR remained unchanged. Suh

et al. (2003) reported a decrease in Grx activity in interfibrillar mitochondria from F344

rat hearts, but no change in Grx activity in subsarcolemmal mitochondria. In F344 rat

liver, aging was reported to have no effect on Grx, TR, or protein disulfide isomerase

activities, while GR activity was increased in females, but not in males (Rikans and

Hornbrook, 1998). In Wistar rat kidney, TR activity was decreased by 50% with aging,

while GR activity remained constant (Santa and Machado, 1986). Finally, Trx content

was reported to increase somewhat with aging in mice, but net Trx activity, and its

association with ASK1, were diminished due to increased S-nitrosylation (Trx-SNO)

(Zhang et al., 2007).

Significance of Bcl-2 and Bcl-xL diminution in susceptibility to cardiomyocyte

apoptosis: Our studies implicate decreases in the anti-apoptotic NFκB target genes Bcl-2

and Bcl-xL in the increased apoptotic susceptibility of Grx1 knock-down cells at rest, and

suggest a special role for Bcl-xL in protection of cardiomyocytes from oxidant-induced

apoptosis. The distinct effects of knocking down Bcl-2 and/or Bcl-xL in H9c2 cells suggest separate, but overlapping, cytoprotective roles for these two proteins in cardiomyocytes, consistent with recent reports documenting both shared (i.e., Bad inhibition, maintenance of mitochondrial homeostasis, and caspase regulation) and distinct (i.e., regulation of calcium homeostasis and antioxidant roles for Bcl-2; DISC inhibition for Bcl-xL) anti-apoptotic mechanisms (Gustafsson and Gottlieb, 2007;

232

Shimizu et al., 1998; Vander Heiden et al., 1999; Krebs et al., 1999; Naumann et al.,

2004; Rong and Distelhorst, 2008; Kane et al., 1993; Voehringer and Meyn, 2000; Hu et al., 1998; Wang et al., 2004). The increased sensitivity of H9c2 cells to Bcl-xL knock-

down compared to Bcl-2 knock-down is consistent with the findings of Feibig et al.

(2006), who observed that Bcl-xL is a more potent anti-apoptotic molecule than Bcl-2 in

T-cells subjected to oxidative stress by Doxorubicin treatment.

4.5 Conclusion

The potential of the thiol-disulfide oxidoreductase enzymes (thioredoxin and glutaredoxin systems) in defense against oxidative stress with aging has been noted previously (Tanaka et al., 2000). However, there has been little direct information about the Grx and Trx systems in this regard. Sohal’s group provided evidence for a protective role of glutaredoxin and GSH in the aging process. Thus, in two strains of houseflies with different longevities, GSH concentration and Grx activity, and GSSG reductase and catalase activities, were higher in the longer-lived strain, indicating they have a higher capacity for oxidative stress (Sohal et al., 1987). By analogy it is reasonable to expect changes in thiol-disulfide oxidoreductase activity to contribute to the complications of aging in mammals also. The results of the current study support this interpretation.

Thus, we present evidence that diminished content and activity of Grx1 predispose cardiomyocytes to apoptosis by decreasing the transcriptional activity of NFκB, resulting in downregulation of its anti-apoptotic target genes, Bcl-2 and Bcl-xL. Targeted knock- down studies of H9c2 cells suggest that both Bcl-2 and Bcl-xL are cytoprotective in unstressed cardiomyocytes, but that Bcl-xL plays a more important role in protection of cardiomyocytes from oxidant-induced apoptosis. Taken together with observations that

233

Grx1 and NFκB activities are diminished with aging in F344 rat heart, these data provide

a mechanistic explanation for why cardiomyocytes in the hearts of the elderly exhibit

increased susceptibility to apoptosis, and further implicate Bcl-xL as an important anti- apoptotic molecule in the heart.

234

Figure 4.1. Grx1 content and activity in heart tissue cytosol and in isolated

myocytes from F344 rats. A, Grx1 content and total Grx activity in cytosolic fractions

of heart tissue from adult (6-10-mo) and elderly (24-28-mo) F344 rats was determined by

semi-quantitative Western blot analysis in which band densities from cytosolic samples

were compared to a standard curve of known amounts of purified Grx1 protein, within

the linear range of concentration dependence on antigen. Grx1 content in elderly rat

hearts (1.1 ± 0.04 pmol Grx1/mg protein) was normalized to content in 6-mo animals (1.7

± 0.02 pmol Grx1/mg protein defined as 100%). Total Grx activity was determined by

monitoring GSH-dependent release of radiolabel from [35S] BSA-SSG as described in

Materials and Methods and in Chrestensen et al. (2000). Activity in elderly hearts (mean

± SEM = 0.94 ± 0.02 nmol [35S] GSSG released/min/mg protein) was normalized to the

activity observed in adult hearts (mean ± SEM = 1.67 ± 0.1 nmol [35S] GSSG

released/min/mg). Open bars, adult; closed bars, elderly. *, p < 0.001 vs. adult (n = 3 for

each age). B-C, Representative microscopy images of cardiomyocytes isolated from adult

(B) and elderly (C) F344 rats. Myocytes were isolated as described in Materials and

Methods, allowed to adhere to laminin-coated dishes for 1 h, and visualized by phase

contrast microscopy (10X magnification). Inset, 40X magnification showing characteristic striations on the rod-shaped cells. D, Total Grx activity in cardiomyocytes isolated from 6-mo F344 rat hearts. Mean values ± SEM were normalized to the value from 6-mo rats (42.5 ± 5.5 nmol NADPH oxidized (i.e., GSSG produced)/min/mg total protein vs. 24.3 ± 6.8 nmol/min/mg for 24-mo rats). *, p < 0.01 (n = 3-4). David Starke generated data shown in A and D, and Elizabeth Sabens took images shown in B-C.

235

236

Figure 4.2. Grx1 content and total Grx activity are decreased via transfection of

targeted siRNA into H9c2 cells. A, Representative Western blot showing decreased

Grx1 content in H9c2 cells transfected with Grx1-targeted siRNA compared to control

siRNA (see Materials and Methods). B, Quantification of densitometric analysis of 6

Western blots of H9c2 cells transfected with control or Grx1-targeted siRNA (Grx1

content in Grx1 knock-down cells is 52 ± 9% (mean ± SEM) decreased compared to

control cells). *, p < 0.03. C, Total Grx activity in H9c2 cells transfected with control or

Grx1-targeted siRNA. Cell lysates were added to assay mix containing Na/K buffer, pH

7.5 (0.1 M), GSH (0.5 mM final), NADPH (0.2 mM final), and GR (2 U/mL, final).

Cysteine-glutathione mixed disulfide (CSSG, 0.1 mM final) was added to initiate the

reaction, and the rate of NADPH oxidation (reflecting the amount of GSSG formed, see

Materials and Methods) was measured by monitoring A340 for 5 min at 30°C. Grx activity (4.5 ± 0.1 nmol NADPH oxidized/min/mg total protein in control cells; 2.6 ± 0.4

nmol/min/mg in Grx1 knock-down cells) was normalized to the value determined in control cells. Data represent mean ± standard deviation (n = 2, p < 0.03). Analogous

decreases in Grx1 content and total Grx activity were observed when Grx1 was stably

knocked down in H9c2 cells using shRNA (see text in Results).

237

238

Figure 4.3. Grx1 knock-down increases apoptotic susceptibility in H9c2 cells. H9c2

cells transfected with control or Grx1-targeted siRNA (see Materials and Methods) were subjected to H2O2 treatment (400 µM for 5 min followed by 24 h recovery (Han et al.,

2004) or simulated ischemia/reperfusion (3 h substrate depletion, hypoxia, and acidosis

followed by 2 h recovery (Kim et al., 2006)), and apoptosis was assessed by Hoechst

staining. A, H9c2 cells in which Grx1 has been diminished by transient transfection with

siRNA. B, H9c2 cells in which Grx1 has been stably knocked-down using shRNA (see

Materials and Methods). Data are represented as mean ± SEM. *, p < 0.02 compared to

no treatment; †, p < 0.02 compared to control siRNA or shRNA (n = 5-10). Open bars,

no treatment; shaded bars, simulated IR; closed bars, H2O2. Dr. Harish Pai generated

data shown in 4.3B.

239

240

Figure 4.4. Grx1-deficient cells exhibit decreased NFκB transcriptional activity

without significant change in NFκB content. A, Western blot analysis of p50 content

in control and Grx1 knock-down cells. Representative Western blot of lysates from H9c2

cells transfected with control or Grx1 siRNA. Band density was normalized to that of

GAPDH, and relative content of p50 was calculated by ratio of band density in Grx1-

deficient cells to control cells (0.8 ± 0.2, n = 3). B, Western blot analysis of p65 content

in control and Grx1 knock-down cells. Representative Western blot of lysates from H9c2

cells transfected with control or Grx1 siRNA. p65 content in knock-down cells was

calculated relative to control cells as described in A (0.8 ± 0.09, n = 4). C, H9c2 cells

were transfected with control or Grx1-targeted siRNA as described in Materials and

Methods, then co-transfected with NFκB-Luciferase and Renilla plasmids. NFκB- dependent transcriptional activity was determined via a standard Dual-Luciferase assay.

Renilla luciferase activity served as a transfection control, and data are represented as ratios of Firefly/Renilla activity, normalized to the mean ratio observed in control cells

(mean ± SEM = 33.9 ± 7.4). *, p < 0.04 (n = 4). Luciferase assays performed by Dr.

Melissa Shelton.

241

242

Figure 4.5. Effect of IKK inhibitor BMS 345541 on NFκB activity and apoptotic

susceptibility in wild-type H9c2 cells. A, Dose-response of BMS 345541 on NFκB-

dependent transcriptional activity. H9c2 cells were transfected with 5X NFκB-Firefly

Luciferase plasmid for 8 h, treated with various concentrations of BMS 345541 for 15 h,

and collected for determination of Luciferase activity. Percent inhibition of NFκB

activity was calculated relative to activity in untreated cells (mean ± SEM = 1264 ± 99

RLU/mg; n = 4). B, Effect of BMS 345541 treatment on sensitivity of H9c2 cells to

H2O2-induced apoptosis. H9c2 cells treated as described above were subjected to 400

μM H2O2 (or no H2O2) for 5 min, then incubated for 24 h and assayed for apoptosis by

Hoechst staining. Open bars (), no treatment; closed bars (), H2O2. *, p < 0.02 vs. no

BMS treatment (no H2O2); †, p < 0.02 vs. no BMS treatment (with H2O2) (n = 4).

243

244

Figure 4.6. Normalized amounts of Bcl-2 and Bcl-xL mRNA in Grx1 knock-down cells relative to control cells. Real-time PCR analysis was performed on mRNA isolated from control and Grx1 knock-down H9c2 cells as detailed in Materials and

Methods. Data are expressed as ratios of the relative quantities of Bcl-2 and Bcl-xL mRNA measured in Grx1 knock-down cells compared to control cells. Open bars (), control cells; closed bars (), Grx1 knock-down cells. Each n represents at least duplicate determinations from a single Grx1 knock-down experiment. For Bcl-2, data represent mean ± SEM (n = 4); for Bcl-xL, data represent mean ± standard deviation (n =

2). For each experiment, Grx1 knock-down was verified separately by Western blot analysis (see representative Western blot in Figure 4.2A). *, p < 0.01.

245

Figure 4.7. Contents of Bcl-2 and Bcl-xL proteins are decreased in Grx1-deficient

H9c2 cells. Content of Bcl-2 and Bcl-xL, both anti-apoptotic NFκB target genes, were compared in control and Grx1 knock-down cells by Western blot analysis. A,

Representative Western blot showing diminished Bcl-2 in Grx1 knock-down cells. B,

Quantification of 6 Western blots showing Bcl-2 content (normalized to actin, 45 ± 14% decrease in Grx1 knock-down cells), * p < 0.03 vs. control siRNA. C, Representative

Western blot showing diminished Bcl-xL in Grx1 knock-down cells. D, Quantification of 4 Western blots showing Bcl-xL content (normalized to GAPDH), 50 ± 14% decrease in Grx1 knock-down cells, * p < 0.02 vs. control siRNA.

246

247

Figure 4.8. Effect of Bcl-2 and Bcl-xL knock-down on apoptotic susceptibility in

H9c2 cells. A, Content of Bcl-2 was diminished via transfection with targeted siRNA as described in Materials and Methods. Representative Western blot analysis of Bcl-2 content in control and Bcl-2 knock-down H9c2 cells (knock-down = 63 ± 4%, n = 3). B,

Content of Bcl-xLwas diminished via transfection with targeted siRNA as described in

Materials and Methods. Representative Western blot analysis of Bcl-xL in control and

Bcl-xL knock-down H9c2 cells (knock-down = 45 ± 7% n = 6). C, Complete knock- down of either Bcl-2 or Bcl-xL does not affect expression of the other protein. H9c2 cells were treated with sufficient concentrations of siRNA targeted for Bcl-2- or Bcl-xL to reduce the target protein to undetectable levels, and content of each protein was determined by Western blotting. Bcl-2 content in Bcl-xL knock-down cells was 116 ±

13% (mean ± SEM) of the value in control cells (n = 9). Bcl-xL content in Bcl-2 knock- down cells was 97 ± 10% (mean ± SEM) of the value in control cells (n = 4). D, Bcl-2 and Bcl-xL content were both diminished via dual transfection with targeted siRNA.

Representative Western blot showing ~40% simultaneous knock-down of both Bcl-2 and

Bcl-xL in H9c2 cells (Bcl-2 knock-down: 45 ± 3%; Bcl-xL knock-down: 37% ± 6, n =

6). E, Apoptosis in control, Bcl-2 knock-down, Bcl-xL knock-down, and Bcl-2/Bcl-xL

double knock-down cells ± H2O2 treatment (400 μM for 5 min, followed by 24h

recovery, see Materials and Methods). Open bars, no treatment; closed bars, H2O2. Data

represent mean % apoptotic cells ± SEM (n = 3-10). *, p < 0.05 vs. control siRNA

(without H2O2); †, p < 0.05 vs. control siRNA (with H2O2).

248

249

250

Figure 4.9. NFκB-dependent transcriptional activity in cardiomyocytes isolated from adult and elderly F344 rat hearts. Cardiomyocytes were isolated from adult and elderly F344 rat hearts via collagenase digestion of connective tissue as described in

Materials and Methods. Immediately following isolation, cardiomyocytes were co- infected with an adenovirus encoding Renilla luciferase (MOI ~20), and an adenovirus encoding the NFκB promoter followed by firefly Luciferase (MOI ~100). Cells were plated, cultured for 24 h, and collected for analysis of luciferase activities. Ratios of firefly luciferase:Renilla luciferase activities were determined for each preparation of cardiomyocytes, then normalized to the value for adult animals (mean ± SEM = 13.1 ±

3.3). For elderly rats, cardiomyocytes from 2 animals were pooled for each experiment

(i.e., each n number). For adult rats, n = 7; for elderly rats, n = 9. *, p < 0.02. Data generated in collaboration with Dr. Melissa Shelton.

251

CHAPTER 5: CONCLUSIONS AND FUTURE DIRECTIONS

5.1 KINETIC CHARACTERIZATION AND POTENTIAL PHYSIOLOGICAL

ROLES OF GRX2

5.1.1 Conclusions and Remaining Questions

Although initial reports emphasized their differences (Johansson et al., 2004;

Johansson et al., 2007), in fact human glutaredoxins 1 and 2 exhibit key catalytic

similarities, including selectivity for glutathione-containing mixed disulfide substrates, a double displacement catalytic mechanism with high commitment to catalysis and the same rate-determining step, equal enhancement of glutathionylation in the presence of

GS, and efficient and selective turnover by GSH and GR relative to the Trx system.

Also like Grx1 (Srinivasan et al., 1997), enhancement of deglutathionylation by hGrx2

can be attributed to an unusually low pKa of the catalytic cysteine (4.6) and enhancement

of the nucleophilicity of GSH as the second substrate, although the difference in pKa

compared to Grx1 (+1 pH unit) and lower enhancement of GSH nucleophilicity result in

an approximately 10-fold lower specific activity for deglutathionylation (Gallogly et al.,

2008).

We and others (Johansson et al., 2004) observed that catalytic efficiency of human and mouse Grx2 for deglutathionylation is about 10-fold lower than that of Grx1, and our studies with GSH and CSSG substrates indicate that this lower efficiency is explained primarily by decreased Vmax,app with little change in KM,app for either substrate. hGrx2 is

also less proficient than Grx1 at scavenging gluathionyl radicals, and at promoting

252

protein glutathionylation in the presence of GSSG. Although hGrx2 is less active than

Grx1 as a deglutathionylating enzyme, consideration of the concentration of both

enzymes in the mitochondria, as well as published reports of mitochondrial pH, led to the

estimation that total deglutathionylation activities are roughly equivalent across

mitochondrial compartments, so long as Grx2 is in its active form.

Evidence supporting the physiological relevance of deglutathionylation by Grx1

continues to grow (Mieyal et al., 2008). However, the importance of Grx2 as a

significant contributor to intracellular deglutathionylation has not been established. First,

it is not yet known whether the deglutathionylating activity exhibited by Grx2 in vitro

takes place in cells also. Second, physiological substrates for deglutathionylation (or

glutathionylation) by monomeric Grx2 are yet to be identified. Finally, the dynamics of

Grx2 regulation by iron-sulfur cluster formation are not understood. Recent studies of

mammalian Grx2 suggest that the majority of Grx2 in mammalian cells is sequestered in inactive 2Fe2S cluster dimers. That is, recombinant hGrx2 purified in the presence of

GSH is isolated from E. coli as the 2Fe2S dimer coordinated with two molecules of GSH

(Lillig et al., 2005; Johansson et al., 2007); hGrx2 monomers form 2Fe2S cluster dimers in an in vitro reconstitution assay, whereas hGrx1 cannot (Berndt et al., 2007); and 55Fe

co-immunoprecipitates with Grx2 in HeLa cells (Lillig et al., 2005). Because

dimerization inhibits the deglutathionylation activity of hGrx2, it is critical to understand

the regulation of Grx2 dimer integrity and assess the resting and activated states of Grx2

in situ. The following experiments are proposed to address these unanswered questions,

increasing understanding of Grx2’s intracellular roles.

253

5.1.2 Is Grx2 active as a deglutathionylating enzyme in situ in resting cells? How

much does it contribute to total cellular deglutathionylation activity?

Experimental Approach: Standard assays for Grx activity in cellular lysate or

tissue homogenate essentially detect only the contribution of cytosolic Grx1, because the

relative content of mitochondrial Grx1 and Grx2 is much smaller. Thus, determination of

Grx2 activity requires subfractionation of samples to isolate only the fraction containing

Grx2 enzyme. For most cells, this corresponds to mitochondrial matrix (Lonn et al.,

2007; Pai et al., 2007)). Thus, mitochondria should be isolated from cells or tissues in

which Grx2 expression has been verified by Western blotting (e.g., HeLa cells (Lillig et

al., 2005), rat heart or liver (Pai et al., 2007). Furthermore, proteins from the

intermembrane space and matrix fractions can be separated by published procedures (Pai

et al., 2007). Grx activity in each fraction can be measured using the standard radiolabel

assay for GSH-dependent degluathionylation of [3H] BSA-SSG (see Chapter 4), then normalized to total mg protein assayed or volume of the mitochondrial space for comparison of total activity between mitochondrial subcompartments. The observed total activity should also be adjusted to account for the pH of each mitochondrial subcompartment, since the standard assay is conducted at pH 7.5. In the absence of activating or inhibiting factors, we predicted that total deglutathionylation activity in each mitochondrial subcompartment would be roughly equivalent (Gallogly et al., 2008;

Chapter 3); however, based on the estimation that 80% of Grx2 is sequestered in inactive dimers (Lillig et al., 2005), we hypothesize that the Grx activity in the mitochondrial matrix will represent about one-fifth of the activity in the intermembrane space.

254

Alternative outcomes: Pai et al. have demonstrated that Grx1 released from the intermembrane space is fully active and corresponds to the amount of Grx1 detected by semi-quantitative Western blot analysis (Pai et al., 2007). If, in contrast, Grx2 activity is undetectable in the mitochondrial matrix fraction, it will be necessary to determine whether activity is truly absent or simply below the detection limit of the assay. The sensitivity of the radiolabel assay can be increased by performing standard addition of isolated, purified Grx to reaction mixtures as described by Tyler Murphy (Tyler Murphy,

MS thesis, Case Western Reserve University). Undetectable activity following standard addition could be explained by very low content of Grx or by inactivation of the enzyme.

These possibilities may be distinguished by performing semi-quantitative Western blot analysis to determine the amount of Grx in each fraction, then using the known specific activities of the respective pure proteins to calculate how much activity should be present if the enzyme is fully active. A predicted activity below the detection limit of the assay

(which can be determined empirically using purified Grx enzyme) is consistent with low abundance of Grx protein, while a predicted activity above the detection limit of the assay suggests enzyme inhibition, although inactivation during sample processing should

also be controlled for by adding standard Grx1 to one portion of the sample at the

beginning of the isolation procedure.

5.1.3 What regulates the release of Grx2 from 2Fe2S cluster dimers? Does dimer

dissociation correspond to increased deglutathionylation activity as expected?

Treatment of Grx2 dimers with oxidants (aerobic conditions, GSSG) or reductants

(ascorbate, dithionite) leads to their dissocation in vitro, yet they are stable when the

concentration of reduced GSH (as ligand) is maintained (Lillig et al., 2005; Berndt et al.,

255

2007). While these observations may at first seem contradictory (i.e., both oxidizing and reducing agents lead to Grx2 dimer dissociation), they are likely explained by distinct effects of the agents on two different sources of Grx2-2Fe2S cluster stability: iron oxidation state and GSH ligand availability. That is, ascorbate and dithionite (both reducing agents) likely induce dimer dissociation via reduction of Fe3+ to Fe2+, destabilizing the Grx2-2Fe2S cluster. Conversely, GSSG treatment (or aerobic conditions, which can oxidize GSH to GSSG) likely causes dimer dissociation via competition with GSH.

In vivo, the role(s) of Fe3+ reduction and GSH depletion on Grx2-2Fe2S cluster homeostasis are unknown. Initial studies should be designed with the following questions in mind: (1) Which mechanism of regulation is being tested (i.e., Fe3+ redox status or GSH levels)? (2) Is the stimulus physiologically relevant? (3) Can the stimulus be targeted to the mitochondria? Two initial experiments are proposed below which address the roles of Fe3+ oxidation state and GSH content on the stability of Grx2-2Fe2S clusters in situ.

Experimental Approach: To address the role of Fe3+ redox state in regulating the integrity of Grx2-2Fe2S clusters, cells (or isolated mitochondria) could be treated with antimycin A, an inhibitor of Complex III within the electron transport chain. Increased generation of superoxide by the electron transport chain due to Antimycin A treatment has been implicated in the dissocation of FeS centers, presumably via reduction of Fe3+ to

Fe2+, leading to damage of FeS proteins (Sturm et al., 2006). Antimycin A treatment is also relevant to cardiac pathophysiology, since normal function of the mitochondrial electron transport chain is impaired during myocardial ischemia (Becker, 2004). Cells

256

already shown to express immunologically detectable Grx2 (e.g., HeLa cells) would be

cultured in the presence of 55Fe as described by Lillig et al. (2005), then treated with

Antimycin A for varying lengths of time. Following cell collection and lysis, Grx2

dimerization status could be assessed as described by Lillig et al. (2005), i.e., via

immunoprecipitation of Grx2 followed by ELISA to determine protein content and

scintillation counting to detect bound iron. Then, Grx2 activity would be determined as

described above for correlation to its dimerization status. We expect that Antimycin A

treatment would reduce the Fe3+ in Grx2-2Fe2S cluster dimers leading to release of

monomeric Grx2 and resulting in a concomitant loss of 55Fe and increase in activity.

To assess the effects of perturbed GSH homeostasis on Grx2-2Fe2S cluster dimer

integrity, HeLa cells could be treated with agents that increase or deplete mitochondrial

GSH. Biotinylated glutathione ethyl ester (Bio-GEE) readily enters cells via diffusion through the plasma membrane and is sequestered inside after cleavage of the ester groups on the molecule. Moreover, Bio-GEE is reported to accumulate in mitochondria, allowing targeting of the treatment to the subcellular compartment of interest

(Zimmermann et al., 2007). We expect Bio-GEE treatment to increase the proportion of dimerized Grx2 (assuming that the biotin moiety will be accommodated in the dimers as

readily as GSH), corresponding to a diminution in deglutathionylation activity within the

mitochondrial matrix. To determine the effects of GSH depletion on Grx2-2Fe2S cluster

dimer integrity and deglutathionylation activity, we propose treating HeLa cells with an

agent that selectively diminishes mitochondrial GSH, e.g., CDDO-IM (Samudio et al.,

2005) and determining the effects on Grx2 dimerization status and deglutathionylation

257

activity as described above. We expect that depletion of mitochondrial GSH will lead to

increased monomerization of Grx2, accompanied by increased activity.

Alternative outcomes: If Antimycin A treatment does not lead to release of Grx2 monomers, then its dissociative effects on other clusters should be confirmed as positive controls. If they dissociate with Antimycin A treatment, then it is possible that Grx2 dimers are not as sensitive to dissociation as other iron-sulfur-associated proteins; alternatively, Grx2-2Fe2S cluster dimers may be protected from the effects of Antimycin

A, e.g., via sequesteration within a particular region of the mitochondrial matrix. This possibility could be investigated by comparing the effects of Antimycin A on intact cells with the effects on a mixture of Antimycin A, mitochondrial membranes containing the components of the electron transport chain, and isolated, purified Grx2-2Fe2S dimers in vitro.

If increasing mitochondrial [GSH] does not increase the proportion of dimeric Grx2

(or vice versa), then in vitro studies of the regulation of Grx2 by GSH should be interpreted with increased caution. Increased [GSH] leads to Grx2 dimerization in vitro; thus a contradictory observation in situ likely reflects an indirect effect of GSH on Grx2 dimerization that is unique to the mitochondrial matrix, i.e., regulation of a factor that in turn affects the Grx2 monomer:dimer ratio. Still, the physiological relevance of this effect should be investigated further. For example, the monomer:dimer ratio (and Grx2 activity) could be investigated in disease models in which GSH is perturbed (e.g.,

Parkinson’s Disease, multiple sclerosis, Type II diabetes mellitus (Dalle-Donne et al.,

2007)), and significant differences in dimerization status and activity could be pursued as

258

potential contributors to disease. In fact, this approach could be utilized also if the effect of GSH manipulation on Grx2 dimerization and activity does turn out as expected.

If dissociation of Grx2 dimers into monomers does not correlate with increased

deglutathionylation activity, then the structure-function relationships observed in vitro

may not correlate to behavior in situ. That is, an inhibitory factor or molecular event

peculiar to the environment of the mitochondrial matrix may prevent deglutathionylation

activity by Grx2 even after release from the inactive dimer. If monomeric Grx2 is not

active as a deglutathionylating enzyme in intact mitochondria, then further

experimentation should focus on identifying the primary enzymatic role of endogenous

Grx2, with glutathionylation (Gallogly et al., 2008; Chapter 3) and FeS cluster synthesis

(Berndt et al., 2007) representing the most logical candidates for initial studies.

5.1.4 What are the physiological substrates for Grx2?

Experimental approach: Three approaches may be taken to identify endogenous

substrates for Grx2. The first approach focuses on mitochondrial matrix proteins shown

to be glutathionylated or deglutathionylated in vitro by Grx, including mitochondrial

Complex I (Beer et al., 2004; Hurd et al., 2008), Complex II (Chen et al., 2007) and

αKGDH (Nulton-Persson et al., 2003; Applegate et al., 2008). To address whether these

proteins are regulated by Grx2 in situ, we propose establishing a link between Grx2

content and protein-SSG status. That is, the protein of interest (i.e., 75 kDa subunit of

Complex I (Beer et al., 2004; Hurd et al., 2008), αKGDH (Applegate et al., 2008)) would be immunoprecipitated from lysates of cells in which Grx2 is knocked down (via siRNA transfection) or overexpressed (using a viral approach analogous to that used for Grx1

(Song et al., 2002; Shelton et al., 2007)), following incubation with NEM to block free

259

sulfhydryl groups from artifactual glutathionylation. The glutathionylation status can be

determined by running the protein on a gel and probing with anti-GSH antibody. An

alternative approach would be to pre-treat the samples with NEM, reduce protein-SSGs

using purified Grx, label the Grx-reduced (i.e., glutathionylated) proteins with a labeled alkylating agent (e.g., NEM-biotin as described by Lind et al. (2002)), then run the samples on a gel and probe with an appropriate detection reagent (e.g., anti-biotin or streptavidin-linked antibody for NEM-biotin). Changes in glutathionylation status at baseline, or following a treatment shown to increase protein glutathionylation in vitro

(e.g., GSSG for Complex I (Beer et al., 2004)) would support regulation by Grx2 in situ.

Follow-up experiments should then focus on whether glutathionylation or deglutathionylation affects function, which could be addressed by correlating the change in glutathionylation status to activity analyses of the affected enzyme. Site-directed mutagenesis of the cysetine that is subject to glutathionylation, which precludes its glutathionylation, can also be used to test its role in the modulation of enzyme function.

That is, cells expressing wild-type and mutant enzymes are treated with a stimulus that induces or reverses glutathionylation of the protein of interest and correlates with a change in the protein’s function. A regulatory role of glutathionylation is supported when cells expressing the mutant protein do not exhibit a change in protein function following exposure to the stimulus (as shown in earlier studies of NF-1 (Bandyopadhyay et al., 1998) and HIV-1 protease (Davis et al., 1997), and more recent studies of Ras

(Adachi et al., 2004a, Pimentel et al., 2006) and SERCA (Adachi et al., 2004b)).

In the second approach, mitochondrial matrix proteins shown to be regulated by cysteine oxidation serve as candidates for regulation by Grx2. For example, the activity

260

of certain components of the mitochondrial permeability transition pore (e.g., ANT

(Halestrap et al., 1997)) are regulated by oxidation of specific sulfhydryl groups, but a

role for glutathionylation has not been confirmed. The ability of these proteins to be

regulated by glutathionylation can be tested in vitro by treatment of isolated proteins or mitochondrial fractions with GSSG or GSH and diamide, followed by activity analysis.

If glutathionylation affects activity, then the protein can be tested as a substrate for Grx2 in glutathionylation and deglutathionylation assays as described in Chapter 4. If a protein’s activity is regulated by reversible glutathionylation, and Grx2 enhances its glutathionylation or deglutathionylation, then determination of glutathionylation as a regulatory mechanism in situ should proceed as described in Approach 1.

The third approach represents a targeted proteomic search for mitochondrial matrix proteins whose glutathionylation status is regulated by Grx2. Grx2 knock-down or overexpressing cells (see above) are lysed, separated by 2-dimensional gel electrophoresis, and probed with an anti-GSH antibody. Comparison of spot density between blots from wild-type, Grx2 knock-down, and Grx2-overexpressing cells will identify proteins whose glutathionylation status changes with manipulation of Grx2.

Candidate proteins can be identified and their glutathionylation status confirmed by digestion and mass spectrometric analysis. Once identified, candidates for regulation by glutathionylation can be verified by in vitro kinetic studies described above, followed by in situ experiments described in Approach 1.

Alternative approaches: It is not known whether all proteins subject to glutathionylation in the mitochondrial matrix are immunoreactive with the anti-GSH antibody. Thus, for approach 3, a technique similar to that of Lind et al. (2002) may be

261

more sensitive. In this method, cellular lysates or tissue homogenates are treated with

NEM to block free sulfhydryl goups and Grx is added to reduce the glutathionylated

proteins as described above. Samples are then incubated with biotin-NEM, run on 2-D

gels, and probed with an anti-biotin or a streptavidin-linked antibody, and changes in

glutathionylation status can be evaluated as described above.

5.1.5 What could be the physiological role(s) of Grx2-2Fe2S in mitochondria?

As discussed in more detail above, one of the major distinctions between human Grx isoforms is the ability of hGrx2 to form a 2Fe2S cluster. Questions that naturally arise from this observation are posed in sections 5.1.2-5.1.4 and include (1) How much monomeric (active) Grx2 is available for catalysis of deglutathionylation in the presence and absence of oxidative stress? (2) What regulates dissociation of Grx2-2Fe2S clusters?

(3) What are the protein targets of monomeric Grx2?

A broader question of equal importance (but less straightforward to answer) is the following: What is the physiological role of the Grx2-2Fe2S cluster? In other words, why is Grx2 complexed to a 2Fe2S cluster in mitochondria, and what are the physiological functions of the monomer and dimer? Studies of iron-sulfur clusters in bacteria (from which eukaryotic mitochondria evolved) provide some insight. For example, FeS centers in a number of prokaryotic proteins serve as redox switches

(reviewed by Imlay, 2006). For many proteins, reduction of an FeS center is inactivating

(e.g., dehydratases lose activity upon reduction of 4Fe4S clusters by ROS and RNS). For other proteins, reduction of an FeS center activates the protein (e.g., SoxR, a transcription

- factor which is activated through reduction of its 2Fe2S center by O2 ).

262

It is plausible that Grx2 could serve as a redox-activated catalyst of reversible

glutathionylation. Under conditions that reduce Fe3+ (e.g., inhibition of the electron

- transport chain with concomitant production of O2 ), the Grx2-2Fe2S cluster could be

disrupted, leading to monomerization (activation) of the enzyme. Grx2 would then serve

as a glutathionylating or deglutathionylating enzyme, depending on the conditions (i.e.,

glutathionylating in the presence of GS, deglutathionylating when GS levels are very

low or absent). In fact, it is conceivable that under extreme conditions (i.e., high rates of radical production), Grx2 would glutathionylate protein substrates in order to protect cysteine sulfhydryl groups from irreversible oxidation; then, once radical production subsides (but before re-sequestration of Grx2 into 2Fe2S cluster dimers), it would

deglutathionylate the proteins, restoring cysteine residues to the reduced state.

The proposed pattern of Grx2 activation and catalysis within the mitochondrial matrix

could contribute to regulation of mitochondrial Complex II (succinate ubiquinone

reductase (SQR) subunit) by reversible glutathionylation. SQR is glutathionylated under

nonstressed conditions in rat heart, but simulated ischemia-reperfusion results in deglutathionylation and inhibition of electron transfer activity (Chen et al., 2007). It is conceivable that under mild oxidative stress conditions, Grx2 is released from 2Fe2S clusters and deglutathionylates SQR, leading to diminished electron transfer and increased electron leak, initially exacerbating the effects of ROS exposure on mitochondrial homeostasis. Upon accumulation of radicals (e.g., GS), Grx2 would be expected to catalyze glutathionylation of SQR, increasing electron transfer activity and protecting the mitochondria from irreversible damage. While certainly plausible, such a

mechanism of regulation of SQR by Grx2 has not yet been investigated.

263

An alternative (or additional) role for the Grx2-2Fe2S dimer within the mitochondrial matrix would be to serve as a storage form of FeS complexes. Many FeS centers in

- prokaryotic proteins readily dissociate in the presence of low concentrations of O2 ,

- H2O2, or ONOO (Imlay, 2006), and some mammalian FeS centers have been

documented to be ROS-sensitive also (Sturm et al., 2006). It is possible that in addition

to (or instead of) its role as a ROS-activated catalyst of reversible glutathionylation, Grx2

supplies FeS centers to other FeS proteins that have become dissociated following

exposure to ROS.

5.2 Regulation of cardiomyocyte apoptosis by Grx1 via modulation of NFκB-

dependent transcription of Bcl-2 and Blc-xL

5.2.1 Conclusions and Remaining Questions

Aging increases the apoptotic susceptibility of cardiomyocytes at baseline and in

response to oxidative stress in animal models (Kajstura et al., 1996; Nitahara et al., 1998;

Liu et al., 2002). We investigated the potential contribution of diminished Grx1 to the increased apoptotic susceptibility associated with aging in cardiomyocytes. Rat

embryonic cardiomyocytes (H9c2 cells) in which Grx1 was diminished to a similar extent

as observed in the aging F344 rat heart exhibited increased apoptotic susceptibility at

baseline, and in response to two oxidative stimuli (H2O2 and simulated IR). NFκB-

dependent transcription, shown previously to be regulated by Grx1 (see Chapters 2, 4),

was diminished in Grx1 knock-down cells, as was mRNA and protein content of the anti-

apoptotic NFκB target genes Bcl-2 and Bcl-xL. Studies using the selective IKK inhibitor

BMS 345541 (Burke et al., 2003) demonstrated that the diminution of NFκB observed

with Grx1 knock-down was sufficient to sensitize H9c2 cells to apoptosis at baseline, and

264

further in the presence of ROS. Moreover, independent knock-down of Bcl-2 or Bcl-xL to the same extent as observed in Grx1-deficient cells suggested roles for each protein in apoptotic susceptibility at baseline. Knock-down of Bcl-xL recapitulated the proapoptotic effect of Grx1 diminution following an oxidative challenge, suggesting an important cytoprotective role for this protein in oxidant-induced apoptosis in cardiomyocytes.

To test whether the mechanism for regulation of apoptosis by Grx1 in H9c2 cells applies to aging cardiomyocytes (in which Grx1 is naturally decreased), NFκB activity was measured in cardiomyocytes isolated from adult and elderly F344 rats. We observed an age-associated decrease in NFκB activity, suggesting correspondence between the in vitro cellular model and the in vivo animal model with respect to mechanisms of apoptotic regulation by Grx1. Additional confirmatory studies in cardiomyocytes from adult and elderly F344 rats are proposed below, and include (1) verification of diminished

Bcl-2 and Bcl-xL content with aging (section 5.2.3) and (2) “rescue” of the apoptotic phenotype of elderly cardiomyocytes by restoration of Grx1 and/or Bcl-xL (section

5.2.4). But perhaps the most intriguing question to guide future studies is what is the mechanism by which Grx1 regulates NFκB in cardiomyocytes? (see next).

5.2.2 What is the site of regulation of the NFκB pathway by Grx1?

Rationale: Multiple lines of evidence indicate that Grx1 regulates NFκB activity in cardiomyocytes, including that Grx1 knock-down in H9c2 cells leads to diminished

NFκB-dependent transcriptional activity and decreased mRNA and protein content of

NFκB target genes; and that diminished NFκB activity is correlated with decreased Grx1

265

in cardiomyocytes isolated from elderly F344 rats (see Chapter 4). However, the

mechanism by which Grx1 modulates NFκB transcriptional activity is unknown.

As discussed above (Chapters 2, 4), numerous proteins in the NFκB signal

transduction pathway have been shown to be regulated by S-glutathionylation in vitro and in situ, including the NFκB transcription factor itself, upstream activators and inhibitors, and regulators of component degradation (reviewed in Figure 2.1, Shelton et al., 2008).

Based on these observations, and that Grx1 is the primary intracellular deglutathionylase

(Chrestensen et al., 2000), it is logical that Grx1 is regulating NFB activity by deglutathionylation of some pathway component or regulator. Because there are many potential sites of regulation by Grx1, priority should be placed on identifying the most likely gluatathionylation target(s) for analysis by analogy to reported findings in other contexts. Then experiments would be designed to test whether regulation of the candidate protein(s) by S-glutathionylation results in sufficient functional changes to account for the apoptotic phenotype observed in Grx1 KD cells.

To be consistent with the observation that decreased Grx1 led to decreased NFκB activity, the glutathionylation target must either activate the NFκB pathway and be inhibited by glutathionylation, or inhibit the pathway and be activated by glutathionylation (assuming that Grx1 is acting as a deglutathionylating enzyme, which is likely in resting cells, see Chapter 1). These criteria eliminate candidates Ras and PTEN, because Ras is activated by S-glutathionylation (Adachi et al., 2004a) and PTEN inhibits

NFkB activity (Mayo et al., 2002; Koul et al., 2001). More specifically, diminution of

Grx1 would be expected to lead to increased levels of Ras-SSG, increasing Ras activity and activating NFκB. For PTEN, the increase in glutathionylation status resulting from

266

diminished Grx1 would be expected to inhibit the enzyme, interfering with its ability to

block NFκB and ultimately leading to increased NFκB activity.

Of the remaining NFκB pathway components regulated by S-glutathionylation, the one best characterized in vitro and in situ is IκB kinase (IKK). IKK phosphorylates the inhibitor of κB protein (IκB), resulting in its ubiquitination, degradation, and concomitant release of the NFκB dimer p50-p65 for translocation to the nucleus (reviewed by Jones

(2003)). IKK is glutathionylated at baseline in retinal Müller cells (Shelton et al., 2009), and reversal of its glutathionylation by Grx1 in lung cells increases NFκB transcriptional activity (Reynaert et al., 2006). Moreover, IKK appears to regulate apoptotic signaling in cardiomyocytes (Craig et al., 2000; Dhingra et al., 2009). Thus, we consider it the most likely target for regulation by Grx1 in cardiomyocytes.

Hypothesis and Experimental Approach: We propose that IKK glutathionylation is increased in Grx1 knock-down cells (or cardiomyocytes from elderly animals in which

Grx1 is diminished), leading to decreased activation of NFκB, decreased transcription of anti-apoptotic NFκB targets, and increased susceptibility to apoptosis. The glutathionylation status of IKK can be assessed by first immunoprecipitating the protein from control and Grx1 knock-down cells (or cardiomyocytes from adult and elderly F344 rats) as described by Shelton et al. (2009). Next, IKK can be released from the

immunoprecipitating antibody and the glutathionylation status of IKK can be analyzed by

the modified biotin-switch assay described above. To determine whether the extent of

glutathionylation of IKK observed with Grx1 deficiency is sufficient to decrease its

activity, purified, recombinant IKK can be glutathionylated in vitro (either using GSSG

or GSH and diamide) and its activity assessed using a standard activity assay using GST-

267

IκBα and 32P-labeled ATP as substrates (Reynaert et al., 2006). If the difference in IKK

glutathionylation observed with Grx1 knock-down results in decreased IKK activity in

vitro, then it is likely that the increase in IKK-SSG affects kinase activity in cells also.

To confirm that IKK inhibition contributes to the apoptotic phenotype of Grx1 knock-

down cells, its activity could be restored selectively by transfecting Grx1 knock-down

cells with plasmid DNA (or virus) encoding IKK. Ideally, transfection conditions would

be optimized to achieve the same level of IKK activity as observed in control cells (i.e.,

non-targeted siRNA, resulting in no Grx1 knock-down). Then, the following would be

compared in control cells, Grx1 knock-down cells transfected with empty vector (i.e., no

IKK), and Grx1 knock-down cells transfected with IKK (i.e., “rescued”): NFκB activity

(via Luciferase assay), relative content of Bcl-2 and Bcl-xL mRNA and protein, and susceptibility to apoptosis with and without H2O2. Rescue of diminished NFκB activity,

decreased Bcl-2 and Bcl-xL, and increased apoptosis would implicate IKK

glutathionylation as a contributory mechanism to the apoptotic phenotype of Grx1 knock-

down cells.

Alternative outcomes and approaches: Increased glutathionylation of IKK would not

explain the increased apoptotic susceptibility of Grx1 knock-down cells if IKK

glutathionylation or activity is not affected by Grx1 knock-down, or if IKK rescue does

not prevent increased apoptotic susceptibility of Grx1 knock-down cells. To identify the

next most likely candidate(s) for regulation of NFκB activity by Grx1, we suggest

identifying the step in NFκB activation that is blocked in Grx1 knock-down cells (e.g.,

nuclear translocation, DNA binding, proteasome activity) to identify potential control

268

points for targeted studies of glutathionylation status (and rescue experiments) as

described above for IKK.

For example, the effect of Grx1 knock-down on nuclear translocation could be

addressed by measuring p50 and p65 content in the cytosol and nucleus via semi-

quantitative Western blotting. If nuclear translocation is diminshed, then the site of

regulation is likely to be at or upstream of translocation, and subsequent studies would focus on relevant pathway components. If there is no difference in NFκB nuclear translocation in Grx1-deficient cells, then blockade of a downstream event, such as DNA

binding, could be addressed by EMSA analysis. Decreased DNA binding with Grx1

knock-down would implicate glutathionylation of p50 and/or p65 as targets of

glutathionylation, since glutathionylation of either protein blocks DNA binding in vitro

(Pineda-Molina et al., 2001; Qanungo et al., 2007).

Following identification of the NFκB-activating event affected by Grx1 knock-down,

the glutathionylation status of the relevant pathway component(s) could be investigated

as described above for IKK. If no protein within the classical NFκB pathway appears to

be regulated by S-glutathionylation, then the possibility of indirect regulation should be

considered, including (1) deglutathionylation of an upstream activator (e.g., Akt, see

below), or (2) transcriptional regulation of an NFκB mediator (see Table 5.1. p. 282, and

below).

Although it has not been shown to be regulated by glutathionylation per se, Akt (an activator of NFκB, (Burow et al., 2000; Dan et al., 2008) exhibits increased anti- apoptotic activity in Grx1-overexpressing H9c2 cells (Murata et al., 2003), confers cytoprotective effects in heart tissue (Miyamoto et al., 2009), and represents a potential

269

mediator of NFκB regulation by Grx1. To determine whether Akt activity is decreased

with Grx1 knock-down, Akt activity could be measured in control and Grx1 knock-down cells as described by Murata and coworkers (2003). If Akt activity is decreased with

Grx1 knock-down, then the contribution of Akt inhibition to apoptotic susceptibility could be tested by treating wild-type H9c2 cells with an Akt inhibitor to achieve the level of decreased activity observed with Grx1 knock-down and determining sensitivity to

H2O2-induced apoptosis. If Akt inhibition is sufficient to increase apoptotic

susceptibility, then subsequent experiments should focus on its role in regulating NFκB- dependent transcription and on the mechanism of regulation by Grx1 (i.e., deglutathionylation, protein-protein interactions, etc.). As for IKK, any mechanistic studies performed in H9c2 cells should be tested also in cardiomyocytes from adult and elderly F344 rats, to confirm the validity of the model system.

An alternative (or additional) mechanism by which Grx1 may regulate NFκB activity is via transcriptional regulation of factors that activate or inhibit the pathway. Indeed, preliminary PCR array analysis (performed as described in Chapter 4, Materials and

Methods) suggests that Grx1 knock-down leads to a decrease in the relative contents of two NFκB activators (CARD10 (Wang et al., 2001c) and Gadd45α (Hoffman and

Liebermann, 2009)) and one NFκB inhibitor (IL-10, reviewed in (Moore et al., 2001)

(Table 5.1, p. 282). The mechanism(s) by which Grx1 regulates the transcription of these genes may involve deglutathionylation of transcription factors or transcriptional regulators, protein-protein interactions, or an as-yet unidentified mechanism of regulation. Before exploring these potential mechanisms of regulation, it would be prudent to first validate changes in protein content following Grx1 knock-down; then

270

confirm that the changes are sufficient to affect NFκB activity. As for other targets of regulation by Grx1, it will also be important to compare the content of the proteins of interest in adult and elderly cardiomyocytes, to provide insight into the relevance of these observations in the aging heart.

5.2.3 Determine relative levels of Bcl-2 and Bcl-xL in adult vs. elderly F344 rats

Rationale: Diminution of Grx1 in H9c2 cells led to decreased NFκB activity as well as decreased mRNA and protein content of anti-apoptotic target genes Bcl-2 and Bcl-xL

(Chapter 4). Primary cardiomyocytes from F344 rats also exhibit decreased Grx1 and

diminished NFκB activity with aging, but age-related differences in Bcl-2 and Bcl-xL

have not been studied. We expect that Bcl-2 and Bcl-xL will be diminished with aging in primary cardiomyocytes from F344 rats because they exhibit decreased Grx1 content and activity, and decreased NFκB-dependent transcription (Chapter 4). Currently, the status of Bcl-2 content in the aging rat heart is unresolved, with reports of no difference in

Bcl-2 mRNA with aging (Liu et al., 2002), a trend towards a decrease in Bcl-2 protein

(Kwak et al., 2006), and a significant diminution in Bcl-2 content (Phaneuf and

Leeuwenburgh, 2002). To our knowledge, there are no published comparisons of Bcl-xL content in adult and elderly cardiomyocytes.

Experimental Approach: mRNA levels of Bcl-2 and Bcl-xL in adult and elderly F344 rat heart tissue can be compared by extracting mRNA from the tissue homogenate using

Trizol (as described in the manufacturer’s protocol), then preparing cDNA and performing real-time PCR analysis using primers for Bcl-2 or Bcl-xL as described in

Chapter 3 (Materials and Methods). Bcl-2 and Bcl-xL content can be determined via

Western blotting in adult vs. elderly F344 rat heart tissue, and in isolated cardiomyocytes

271

(for correlation with measurements of decreased NFκB activity in isolated

cardiomyocytes) as described in Chapter 4 (Materials and Methods). Before performing

the direct comparison, it is important to determine the linear range of detection for each

antibody by generating standard curves of tissue homogenate or cell lysate and probing

for the protein of interest. It is also important to confirm (by Western blotting or by

activity analysis) that Grx1 is decreased with aging in the samples analyzed for Bcl-2 and

Bcl-xL.

Alternative outcomes: We have shown that selective diminution of Grx1 in H9c2

cells to a similar extent as observed in aging cardiomyocytes leads to decreased activity

of NFκB, a transcription factor that regulates Bcl-2 and Bcl-xL transcription, as well as

diminution of Bcl-2 and Bcl-xL by approximately one-half (Chapter 4). However, aging

is a complex phenotype (see Juhaszova et al., 2005) in which Grx1 diminution represents

one of many molecular changes; hence, it is possible that cardiomyocytes from aging rats

do not exhibit diminished content Bcl-2 and Bcl-xL, even if Grx1 and NFκB activity are

both decreased (see Chapter 4).

For example, many transcription factors besides NFκB are reported to regulate the

transcription of Bcl-2 and Bcl-xL (Hewitt et al., 1995; Li et al., 2003; Kobayashi et al.,

2006; Sakamoto and Frank, 2009; Connors et al., 2009; Kuo and Chang, 2007; Sevilla et

al., 2001). It is possible that aging results in activation or inhibition of one or more of

these transcriptional regulators, nullifying or even overcoming the effects of decreased

NFκB activity on Bcl-2 and Bcl-xL mRNA content. Because of the large number of

transcriptional regulators identified for Bcl-2 and Bcl-xL, systematic analysis of each candidate transcription factor’s activity would be impractical. However, the most likely

272

candidates could be identified by considering which transcription factors regulate both

Bcl-2 and Bcl-xL (e.g., GATA4, Kobayashi et al., 2006; Kuo et al., 2007), as well as

those shown to be active in cardiomyocytes.

If mRNA levels of Bcl-2 and Bcl-xL are decreased with aging, but protein levels are

not, this may reflect an age-related slowing of the rate of protein degradation. Multiple

studies suggest an overall slowdown of protein degradation with aging (reviewed in

(Martinez-Vicente et al., 2005), which could be confirmed by comparing activities of the

implicated pathways (e.g., ubiquitin-proteasome system, lysosomal degradation) in

cardiomyocytes isolated from adult and elderly F344 rats.

If Bcl-2 and Bcl-xL are not diminished with aging in the F344 rat heart, then there are

likely to be different (or additional) mechanisms for increased apoptotic susceptibility in

the aging heart compared to Grx1 knock-down cells. One strategy for identifying such

mechanisms is to perform quantitative PCR array analysis focused on regulators of

apoptosis using samples from adult and elderly F344 rat heart, and control and Grx1

knock-down H9c2 cells. In these real-time PCR experiments, relative quantities of 84

apoptosis-related genes are compared in up to 4 samples. Pilot studies in H9c2 cells

identified 8 genes that are increased or decreased substantially with Grx1 knock-down

(Table 5.1, p. 282). Comparison of these changes to the changes observed with aging

would provide insight into the correspondence between Grx1 knock-down cells and

cardiomyocytes from aging animals with regard to one aspect of apoptotic regulation

(i.e., transcriptional regulation of apoptotic mediators).

273

5.2.4 Can elderly cardiomyocytes be “rescued” from their apoptotic phenotype by

addition of Grx1 or Bcl-xL?

Rationale: Mechanistic studies in H9c2 cells indicate that diminution of Bcl-xL in

Grx1 knock-down cells accounts for their increased susceptibility to apoptosis (Chapter

4). Thus, it is logical that restoration of of Bcl-xL to levels observed in wild-type H9c2

cells will “rescue” the apoptotic phenotype, decreasing apoptosis to levels observed in

cells transfected with control siRNA. If Bcl-xL is also decreased in aging

cardiomyocytes (see 5.2.2), then its restoration should improve survival at baseline and

following oxidative insult. Thus, rescuing Grx1-deficient cardiomyocytes by restoring

Bcl-xL would confirm the proposed mechanism by which Grx1 regulates apoptotic

susceptibility in cardiomyocytes (Scheme 2.1, p. 148). Moreover, the ability to rescue aging cardiomyocytes via restoration of Bcl-xL or Grx1 would provide a foundation for the development of therapeutic agents designed to minimize the adverse effects of aging on cardiovascular health.

Hypothesis and Experimental Approach: Because cardiomyocytes from aging F344 rats exhibit decreased Grx1 content and activity, decreased NFκB activity (Chapter 4), and increased apoptosis (Kajstura et al., 1996; Nitahara et al., 1998; Liu et al., 2002), we

hypothesize that restoration of Grx1 content to the level observed in adult

cardiomyocytes will improve survival at baseline and following an oxidative challenge.

Based on observations that Bcl-xL knockdown sensitizes H9c2 cells to apoptosis

(Chapter 4), and that Bcl-xL overexpression decreases apoptosis in rodent heart (Huang

et al., 2003), we expect restoration of Bcl-xL in Grx1-deficient cells to decrease apoptotic susceptibility to levels observed in control cells (i.e., no Grx1 knock-down for H9c2

274

cells, adult cardiomyocytes for F344 rats). Grx1 and Bcl-xL levels can be restored in

H9c2 cells with Grx1 knock-down via transfection of a plasmid encoding rat Grx1 or

Bcl-xL; alternatively, a viral method of gene delivery can be utilized. It is important to

increase Grx1/Bcl-xL content to match that observed in wild-type cells so that the

experiment tests the effect of Grx1/Bcl-xL restoration rather than overexpression, which

has already been shown to increase cardiomyocyte viability following oxidative insult

(Huang et al., 2003). Next, apoptosis should be assessed at baseline (i.e., no stimulus)

and following an oxidative insult previously shown to trigger apoptosis (i.e., H2O2

treatment as described in Chapter 4). The effect of Grx1/Bcl-xL “rescue” will be

determined by comparing the amount of apoptosis observed in Grx1 knock-down cells

with augmented Grx1/Bcl-xL to the amount in Grx1 knock-down cells transfected with

empty vector, and to control cells (i.e., no Grx1 knock-down, normal Bcl-xL). A similar approach can be utilized for cardiomyocytes from elderly F344 rats; however, Grx1/Bcl- xL must be restored using viral delivery since primary cardiomyocytes are resistant to transfection by plasmid DNA (Djurovic et al., 2004). Indeed, infection of cardiomyocytes from 6-mo F344 rats with adenovirus encoding Grx1 resulted in increased Grx1 expression (experiments performed by Dr. Melissa Shelton), suggesting that this approach may be feasible for cardiomyocytes from 24-mo animals as well.

Alternative outcomes: If restoration of Grx1 rescues Grx1 knock-down cells, but not elderly cardiomyocytes, from increased apoptotic susceptibility, then it is likely that diminution of Grx1 is not the only molecular change that contributes to increased apoptosis in elderly cardiomyocytes, and therapies aimed at restoring Grx1 to adult levels would provide little benefit to the elderly. However, it is possible that Grx1

275

overexpression could confer a therapeutic benefit regardless of age, since Grx1

overexpression in H9c2 cells increased their tolerance to H2O2 treatment and Grx1

transgenic mice exhibit improved post-MI cardiac function in comparison to control

animals (Malik et al., 2008). Moreover, acute overexpression of Grx1 (i.e., within the

context of gene therapy) may confer an even greater benefit, since there is less of an

opportunity to activate compensatory mechanisms than in an embryonic transgenic

model.

If Grx1 restoration does not rescue the apoptotic phenotype in Grx1 knock-down

H9c2 cells or elderly cardiomyocytes, then it is possible that the combination of siRNA

treatment (for knock-down) and subsequent transfection (for “rescue”) represents a

cellular stress that cannot be overcome by restoration of Grx1 levels. If this is suspected,

then cellular toxicity could be assessed by determining the apoptotic susceptibility of

Grx1 knock-down cells with and without transfection of the empty vector used as a

control treatment in rescue experiments. If the second transfection siginificantly

increases the apoptotic susceptibility of Grx1 knock-down cells, then alternative (less

toxic) methods of Grx1 (or Bcl-xL) “rescue” should be considered. Potential approaches

to decreasing toxicity include varying the method of rescue (e.g., viral infection vs.

transfection of a plasmid), the transfection agents used (e.g., Lipofectamine 2000 vs.

Lipofectamine), and/or the transfection conditions (e.g., dose, incubation time, % serum

in the transfection medium).

An alternative to transfection for restoration of Grx1 or Bcl-xL levels is treatment of

Grx1 KD cells with either protein fused to HIV-1 TAT. The HIV-1 TAT protein contains an 11 amino acid protein transduction domain that, when fused to a protein, results in

276

rapid and receptor-independent entry of the tagged protein into the cytosol (Cao et al.,

2002; Hotchkiss et al., 2006). Studies in neurons (Hotchkiss et al., 2006) and

lymphocytes (Cao et al., 2002) indicate that TAT-tagged Bcl-xL permeates cell membranes and confers anti-apoptotic effects in the presence of ischemia and sepsis, respectively. In the context of ischemia-reperfusion (IR), TAT-mediated delivery of the

Bcl-xL BH4 domain protected isolated rat hearts from post-IR apoptosis (Ono et al.,

2005); however, the specificity of this effect for Bcl-xL is difficult to determine because its BH4 domain is shared with Bcl-2. Since our findings suggested distinct anti-apoptotic effects of Bcl-2 and Bcl-xL in cardiomyocytes subjected to oxidative stress (see Chapter

4), we recommend tagging and utilizing the entire Bcl-xL protein for rescue experiments in Grx1 knock-down cells. Advantages to the TAT-tagging method of rescue include a possible reduction in toxicity compared to sequential transfections, as well as the potential to facilitate dose-response studies. Although the TAT-tagged Grx1 and/or Bcl- xL proteins would have to be synthesized and purified in-house, their use represents a specific (and potentially less toxic) approach to rescue experiments.

If Grx1 restoration rescues the apoptotic phenotype of Grx1 knock-down H9c2 cells and elderly cardiomyocytes, but Bcl-xL does not, then it is possible that the knock-down of Bcl-xL by siRNA described in Chapter 4 was not selective, and an unrelated molecular change was responsible for the increased apoptosis in Bcl-xL knock-down cells. If Bcl- xL restoration rescues Grx1 knock-down cells, but not elderly cardiomyocytes, from increased apoptosis, then it is likely that age-related changes in addition to Bcl-xL diminution contribute to the increased apoptotic susceptibility of these cells, and that rescue of Bcl-xL alone is not sufficient to restore them to the adult phenotype. As above,

277

Bcl-xL overexpression may still confer a benefit to elderly cardiomyocytes, since Bcl-xL

transgenic animals exhibit increased resistance to MI-induced cardiac injury (Huang et

al., 2003).

Alternative approach:

A logical alternative to restoring Grx1 levels to the aging heart is preventing the initial loss in Grx1 protein. To date, the reason for the age-related decline in Grx1 is unknown, but may result from decreased transcription or induction, increased degradation, reduced stability, or a combination. Exploring the basis for the decrease in

Grx1 protein and activity with aging would begin by investigating age-related changes in mRNA content and protein half-life; then, subsequent experiments would focus on identifying reasons for the decreased transcription or increased degradation. At present, mechanisms of regulation of Grx1 content are not well understood, but hormonal, chemical, and metabolic stimuli have been implicated in various cell types (see Section

1.3.2A; Gallogly et al., 2009). Notably, a recent study in rats has linked broccoli ingestion with upregulation of various TDOR enzymes, including Grx1 (Muhkerjee et al.,

2008). Elucidation of the mechanism by which Grx1 levels decline with old age would increase understanding of the mechanisms by which Grx1 is regulated in situ, and identify potential strategies to prevent its decline with aging, an approach that has the potential to be less traumatic and more cost-effective than intensive medical treatment

(especially if the intervention is a simple dietary modification or administration of a selective inducing agent).

278

5.2.5 Potential role for Grx1 and/or Bcl-xL as therapeutic targets in the aging heart

Several studies demonstrate cytoprotective roles for Grx1 and/or Bcl-xL in cardiomyocytes in vitro and in vivo (Murata et al., 2003; Huang et al., 2003), and the

studies presented in Chapter 4 suggest a potential mechanism for their protective effects.

These observations naturally lead to the question of whether Grx1 and Bcl-xL represent

therapeutic agents in heart disease, particularly for elderly patients.

To date, no selective activators or inhibitors have been identified for Grx1, so the

only potential mechanism for its restoration is gene therapy. A considerable risk

associated with this approach is lack of selectivity, since Grx1 regulates so many cellular

functions (e.g., inflammation, metabolism, calcium homeostasis, see Mieyal et al., 2008).

This risk would be compounded by the possibility that gene delivery, if not precisely

titrated, could result in overexpression of Grx1 compared to adult levels (vs. rescue to

match the content observed in adults). This possibility warrants particular consideration

in the context of NFκB regulation by Grx1, because excessive activation of this pathway,

while beneficial for cellular survival, could also lead to deleterious inflammation, a

condition already suspected to exacerbate age-related pathologies (Adler et al., 2007).

Depending on results of experiments described in section 5.2.3, restoration of Bcl-xL

could represent a more selective approach to target increased apoptotic susceptibility with

aging. Indeed, therapeutic agents that perform one function of Bcl-xL (i.e., caspase

inhibitors) minimize post-MI cardiac injury in rats (reviewed in (Ogata and Takahashi,

2003), and in contrast to Grx1, the overexpression of Bcl-xL seems less likely to be

associated with deleterious non-specific effects. Restoration of Bcl-xL activity could be

achieved through gene therapy, targeted delivery of Bcl-xL protein (as described by Chen

279

et al., 2002), or the development of Bcl-xL activators/stabilizers, and represents an exciting frontier in the field of cardiovascular medicine.

280

Table 5.1. Apoptotic regulators exhibiting substantial changes in mRNA content with Grx1 knock-down. Relative quantities of mRNA were measured in control and

Grx1 knock-down cells by real-time PCR analysis using a PCR array plate pre-loaded with primers for 84 apoptosis-related genes (described in Chapter 4, Materials and

Methods). Samples from two different Grx1 knock-down experiments were measured, and each sample was measured in duplicate. For each experiment, diminution of Grx1 knock-down was verified by Western blotting prior to real-time PCR analysis. Relative quantities of mRNA were normalized to GAPDH, then to the value in control cells. Data are represented as the mean of two determinations ± standard deviation. When mRNA content was not detectable in one of two determinations, the data was not included. Any gene increased by≥ 2 -fold or decreased by at least 50% with Grx1 knock-down was considered to change substantially.

281

Fold change with Gene Grx1 knock-down Function/Reference Card_10 ↓ 0.7 ± 0.1 Caspase recruitment domain family, member 10; predicted proposed to form a scaffold for assembly of a Bcl-10 signaling complex that activates NFκB (Wang et al., 2001c)

Caspase ↑ 4.2 ± 1.4 Terminal differentiation, especially in 14 keratinocytes (Denecker et al., 2008)

Casp8ap2 ↑ 2.2 ± 0.2 Caspase 8-associated protein 2 (FLASH); activates predicted caspase-8 and facilitates Fas-induced apoptosis (Flotho et al., 2006)

Dffb ↓ 0.6 ± 0.03 DNA fragmentation factor, beta subunit, or caspase- activated DNAse (CAD); DNase activated by caspases 3 and 7 to facilitate apoptosis (Neimanis et al., 2007)

Gadd45a ↓ 0.5 ± 0.03 Growth arrest and DNA damage-inducible 45, alpha; stress signaling, hematopoiesis, innate immunity, NFκB activation and JNK inhibition (Hoffman and Libermann, 2009)

IL-10 ↑ 5.8 ± 4.5 Cytokine synthesis inhibitory factor (CSIF); differentiation and function of regulator T cells (Moore et al., 2001), can block NFκB activity

Mapk8ip ↓ 0.4 ± 0.01 Mitogen activated protein kinase 8 interacting protein; scaffold for JNK stress signaling pathway components (Jaeschke et al., 2004)

Trp73 ↑ 4.0 ± 2.0 Transformation-related protein 73; 2 isoforms confer predicted opposing effects on apoptosis (Tomasini et al., 2008)

282

BIBLIOGRAPHY

Abraham,R.T., Benson,L.M., and Jardine,I. (1983). Synthesis and pH-dependent stability of purine-6-sulfenic acid, a putative reactive metabolite of 6-thiopurine. J. Med. Chem. 26, 1523-1526.

Adachi,T., Pimentel,D.R., Heibeck,T., Hou,X., Lee,Y.J., Jiang,B., Ido,Y., and Cohen,R.A. (2004a). S-glutathiolation of Ras mediates redox-sensitive signaling by angiotensin II in vascular smooth muscle cells. J. Biol. Chem. 279, 29857-29862.

Adachi,T., Weisbrod,R.M., Pimentel,D.R., Ying,J., Sharov,V.S., Schoneich,C., and Cohen,R.A. (2004b). S-Glutathiolation by peroxynitrite activates SERCA during arterial relaxation by nitric oxide. Nat. Med. 10, 1200-1207.

Adler,A.S., Sinha,S., Kawahara,T.L., Zhang,J.Y., Segal,E., and Chang,H.Y. (2007). Motif module map reveals enforcement of aging by continual NF-kappaB activity. Genes Dev. 21, 3244-3257.

Anathy,V., Aesif,S.W., Guala,A.S., Havermans,M., Reynaert,N.L., Ho,Y.S., Budd,R.C., and Janssen-Heininger,Y.M. (2009). Redox amplification of apoptosis by caspase- dependent cleavage of glutaredoxin 1 and S-glutathionylation of Fas. J. Cell Biol. 184, 241-252.

Applegate,M.A., Humphries,K.M., and Szweda,L.I. (2008). Reversible inhibition of alpha-ketoglutarate dehydrogenase by hydrogen peroxide: glutathionylation and protection of lipoic acid. Biochemistry 47, 473-478.

Aracena-Parks,P., Goonasekera,S.A., Gilman,C.P., Dirksen,R.T., Hidalgo,C., and Hamilton,S.L. (2006). Identification of cysteines involved in S-nitrosylation, S- glutathionylation, and oxidation to disulfides in ryanodine receptor type 1. J. Biol. Chem. 281, 40354-40368.

Arnelle,D.R. and Stamler,J.S. (1995). NO+, NO, and NO- donation by S-nitrosothiols: implications for regulation of physiological functions by S-nitrosylation and acceleration of disulfide formation. Arch. Biochem. Biophys. 318, 279-285.

Ashton,K.J., Willems,L., Holmgren,K., Ferreira,L., and Headrick,J.P. (2006). Age- associated shifts in cardiac gene transcription and transcriptional responses to ischemic stress. Exp. Gerontol. 41, 189-204.

Aslund,F., Ehn,B., Miranda-Vizuete,A., Pueyo,C., and Holmgren,A. (1994). Two additional glutaredoxins exist in Escherichia coli: glutaredoxin 3 is a hydrogen donor for ribonucleotide reductase in a thioredoxin/glutaredoxin 1 double mutant. Proc. Natl. Acad. Sci. U. S. A 91, 9813-9817.

283

Aykac-Toker,G., Bulgurcuoglu,S., and Kocak-Toker,N. (2001). Effect of peroxynitrite on glutaredoxin. Hum. Exp. Toxicol. 20, 373-376.

Baker,A.F., Landowski,T., Dorr,R., Tate,W.R., Gard,J.M., Tavenner,B.E., Dragovich,T., Coon,A., and Powis,G. (2007). The antitumor agent imexon activates antioxidant gene expression: evidence for an oxidative stress response. Clin. Cancer Res. 13, 3388-3394.

Bandyopadhyay,S., Starke,D.W., Mieyal,J.J., Gronostajski,R.M. (1998). Thioltransferase (glutaredoxin) reactivates the DNA-binding activity of oxidation- inactivated nuclear factor 1. J. Biol. Chem. 273, 392-397.

Barrett,W.C., DeGnore,J.P., Keng,Y.F., Zhang,Z.Y., Yim,M.B., and Chock,P.B. (1999a). Roles of superoxide radical anion in signal transduction mediated by reversible regulation of protein-tyrosine phosphatase 1B. J. Biol. Chem. 274, 34543-34546.

Barrett,W.C., DeGnore,J.P., Konig,S., Fales,H.M., Keng,Y.F., Zhang,Z.Y., Yim,M.B., and Chock,P.B. (1999b). Regulation of PTP1B via glutathionylation of the active site cysteine 215. Biochemistry 38, 6699-6705.

Becker,L.B. (2004). New concepts in reactive oxygen species and cardiovascular reperfusion physiology. Cardiovasc. Res. 61, 461-470.

Beer,S.M., Taylor,E.R., Brown,S.E., Dahm,C.C., Costa,N.J., Runswick,M.J., and Murphy,M.P. (2004). Glutaredoxin 2 catalyzes the reversible oxidation and glutathionylation of mitochondrial membrane thiol proteins: implications for mitochondrial redox regulation and antioxidant DEFENSE. J. Biol. Chem. 279, 47939- 47951.

Belli,G., Polaina,J., Tamarit,J., De La Torre,M.A., Rodriguez-Manzaneque,M.T., Ros,J., and Herrero,E. (2002). Structure-function analysis of yeast Grx5 monothiol glutaredoxin defines essential amino acids for the function of the protein. J. Biol. Chem. 277, 37590- 37596.

Bergmann,M.W., Loser,P., Dietz,R., and von,H.R. (2001). Effect of NF-kappa B Inhibition on TNF-alpha-induced apoptosis and downstream pathways in cardiomyocytes. J. Mol. Cell Cardiol. 33, 1223-1232.

Berndt,C., Hudemann,C., Hanschmann,E.M., Axelsson,R., Holmgren,A., and Lillig,C.H. (2007). How does iron-sulfur cluster coordination regulate the activity of human glutaredoxin 2? Antioxid. Redox. Signal. 9, 151-157.

Biteau,B., Labarre,J., and Toledano,M.B. (2003). ATP-dependent reduction of cysteine- sulphinic acid by S. cerevisiae sulphiredoxin. Nature 425, 980-984.

Blom,N., Gammeltoft,S., and Brunak,S. (1999). Sequence and structure-based prediction of eukaryotic protein phosphorylation sites. J. Mol. Biol. 294, 1351-1362.

284

Blom,N., Sicheritz-Ponten,T., Gupta,R., Gammeltoft,S., and Brunak,S. (2004). Prediction of post-translational glycosylation and phosphorylation of proteins from the amino acid sequence. Proteomics. 4, 1633-1649.

Bonavita,F., Stefanelli,C., Giordano,E., Columbaro,M., Facchini,A., Bonafe,F., Caldarera,C.M., and Guarnieri,C. (2003). H9c2 cardiac myoblasts undergo apoptosis in a model of ischemia consisting of serum deprivation and hypoxia: inhibition by PMA. FEBS Lett. 536, 85-91.

Bradshaw,T.P., McMillan,D.C., Crouch,R.K., and Jollow,D.J. (1995). Identification of free radicals produced in rat erythrocytes exposed to hemolytic concentrations of phenylhydroxylamine. Free Radic. Biol. Med. 18, 279-285.

Brocheriou,V., Hagege,A.A., Oubenaissa,A., Lambert,M., Mallet,V.O., Duriez,M., Wassef,M., Kahn,A., Menasche,P., and Gilgenkrantz,H. (2000). Cardiac functional improvement by a human Bcl-2 transgene in a mouse model of ischemia/reperfusion injury. J. Gene Med. 2, 326-333.

Burke,J.R., Pattoli,M.A., Gregor,K.R., Brassil,P.J., MacMaster,J.F., McIntyre,K.W., Yang,X., Iotzova,V.S., Clarke,W., Strnad,J., Qiu,Y., and Zusi,F.C. (2003). BMS-345541 is a highly selective inhibitor of I kappa B kinase that binds at an allosteric site of the enzyme and blocks NF-kappa B-dependent transcription in mice. J. Biol. Chem. 278, 1450-1456.

Burow,M.E., Weldon,C.B., Melnik,L.I., Duong,B.N., Collins-Burow,B.M., Beckman,B.S., and McLachlan,J.A. (2000). PI3-K/AKT regulation of NF-kappaB signaling events in suppression of TNF-induced apoptosis. Biochem. Biophys. Res. Commun. 271, 342-345.

Burwell,L.S., Nadtochiy,S.M., Tompkins,A.J., Young,S., and Brookes,P.S. (2006). Direct evidence for S-nitrosation of mitochondrial complex I. Biochem. J. 394, 627-634.

Bushweller,J.H., Aslund,F., Wuthrich,K., and Holmgren,A. (1992). Structural and functional characterization of the mutant Escherichia coli glutaredoxin (C14----S) and its mixed disulfide with glutathione. Biochemistry 31, 9288-9293.

Bushweller,J.H., Billeter,M., Holmgren,A., and Wuthrich,K. (1994). The nuclear magnetic resonance solution structure of the mixed disulfide between Escherichia coli glutaredoxin(C14S) and glutathione. J. Mol. Biol. 235, 1585-1597.

Cadenas,E. (1995). Thiyl radical formation during thiol oxidation by ferrylmyoglobin. Methods Enzymol. 251, 106-116.

Caffrey,J.L., Boluyt,M.O., Younes,A., Barron,B.A., O'Neill,L., Crow,M.T., and Lakatta,E.G. (1994). Aging, cardiac proenkephalin mRNA and enkephalin peptides in the Fisher 344 rat. J. Mol. Cell Cardiol. 26, 701-711.

285

Camaschella,C., Campanella,A., De,F.L., Boschetto,L., Merlini,R., Silvestri,L., Levi,S., and Iolascon,A. (2007). The human counterpart of zebrafish shiraz shows sideroblastic- like microcytic anemia and iron overload. Blood 110, 1353-1358.

Cao,G., Pei,W., Ge, H., Liang,Q., Luo,Y., Sharp,F.R., Lu,A., Ran,R., Graham,S.H., Chen,J. (2002). In Vivo Delivery of a Bcl-xL Fusion Protein Containing the TAT Protein Transduction Domain Protects against Ischemic Brain Injury and Neuronal Apoptosis. J. Neurosci. 22, 5423-5431.

Carlberg,I. and Mannervik,B. (1985). Glutathione reductase. Methods Enzymol. 113, 484-490.

Chandra,A., Srivastava,S., Petrash,J.M., Bhatnagar,A., and Srivastava,S.K. (1997). Modification of aldose reductase by S-nitrosoglutathione. Biochemistry 36, 15801- 15809.

Chao,D.T., Linette,G.P., Boise,L.H., White,L.S., Thompson,C.B., and Korsmeyer,S.J. (1995). Bcl-XL and Bcl-2 repress a common pathway of cell death. J. Exp. Med. 182, 821-828.

Chen,F.C. and Ogut,O. (2006). Decline of contractility during ischemia-reperfusion injury: actin glutathionylation and its effect on allosteric interaction with tropomyosin. Am. J. Physiol Cell Physiol 290, C719-C727.

Chen,M., Won,D.J., Krajewski,S., and Gottlieb,R.A. (2002). Calpain and mitochondria in ischemia/reperfusion injury. J. Biol. Chem. 277, 29181-29186.

Chen,Y.R., Chen,C.L., Pfeiffer,D.R., and Zweier,J.L. (2007). Mitochondrial complex II in the post-ischemic heart: oxidative injury and the role of protein S-glutathionylation. J Biol. Chem. 282, 32640-32654.

Chen,Z., Chua,C.C., Ho,Y.S., Hamdy,R.C., and Chua,B.H. (2001). Overexpression of Bcl-2 attenuates apoptosis and protects against myocardial I/R injury in transgenic mice. Am. J. Physiol Heart Circ. Physiol 280, H2313-H2320.

Chiu,J.J., Usami,S., and Chien,S. (2008). Vascular endothelial responses to altered shear stress: Pathologic implications for atherosclerosis. Ann. Med. 1-10.

Chrestensen,C.A., Eckman,C.B., Starke,D.W., and Mieyal,J.J. (1995). Cloning, expression and characterization of human thioltransferase (glutaredoxin) in E. coli. FEBS Lett. 374, 25-28.

Chrestensen,C.A., Starke,D.W., and Mieyal,J.J. (2000). Acute cadmium exposure inactivates thioltransferase (Glutaredoxin), inhibits intracellular reduction of protein- glutathionyl-mixed disulfides, and initiates apoptosis. J. Biol. Chem. 275, 26556-26565.

286

Claiborne,A., Miller,H., Parsonage,D., and Ross,R.P. (1993). Protein-sulfenic acid stabilization and function in enzyme catalysis and gene regulation. FASEB J. 7, 1483- 1490.

Claiborne,A., Yeh,J.I., Mallett,T.C., Luba,J., Crane,E.J., III, Charrier,V., and Parsonage,D. (1999). Protein-sulfenic acids: diverse roles for an unlikely player in enzyme catalysis and redox regulation. Biochemistry 38, 15407-15416.

Clavreul,N., Adachi,T., Pimental,D.R., Ido,Y., Schoneich,C., and Cohen,R.A. (2006a). S- glutathiolation by peroxynitrite of p21ras at cysteine-118 mediates its direct activation and downstream signaling in endothelial cells. FASEB J. 20, 518-520.

Clavreul,N., Bachschmid,M.M., Hou,X., Shi,C., Idrizovic,A., Ido,Y., Pimentel,D., and Cohen,R.A. (2006b). S-glutathiolation of p21ras by peroxynitrite mediates endothelial insulin resistance caused by oxidized low-density lipoprotein. Arterioscler. Thromb. Vasc. Biol. 26, 2454-2461.

Cohen,N.S., Cheung,C.W., and Raijman,L. (1987). Measurements of mitochondrial volumes are affected by the amount of mitochondria used in the determinations. Biochem. J. 245, 375-379.

Colell,A., Ricci,J.E., Tait,S., Milasta,S., Maurer,U., Bouchier-Hayes,L., Fitzgerald,P., Guio-Carrion,A., Waterhouse,N.J., Li,C.W., Mari,B., Barbry,P., Newmeyer,D.D., Beere,H.M., and Green,D.R. (2007). GAPDH and autophagy preserve survival after apoptotic cytochrome c release in the absence of caspase activation. Cell 129, 983-997.

Connors,S.K., Balusu,R., Kundu,C.N., Jaiswal,A.S., Gairola,C.G., and Narayan,S. (2009). C/EBPbeta-mediated transcriptional regulation of bcl-xl gene expression in human breast epithelial cells in response to cigarette smoke condensate. Oncogene 28, 921-932.

Coppock,D.L. and Thorpe,C. (2006). Multidomain flavin-dependent sulfhydryl oxidases. Antioxid. Redox. Signal. 8, 300-311.

Craig,R., Larkin,A., Mingo,A.M., Thuerauf,D.J., Andrews,C., McDonough,P.M., and Glembotski,C.C. (2000). p38 MAPK and NF-kappa B collaborate to induce interleukin-6 gene expression and release. Evidence for a cytoprotective autocrine signaling pathway in a cardiac myocyte model system. J. Biol. Chem. 275, 23814-23824.

Cruz,C.M., Rinna,A., Forman,H.J., Ventura,A.L., Persechini,P.M., and Ojcius,D.M. (2007). ATP activates a reactive oxygen species-dependent oxidative stress response and secretion of proinflammatory cytokines in macrophages. J. Biol. Chem. 282, 2871-2879.

Dal Monte,M., Cecconi,I., Buono,F., Vilardo,P.G., Del Corso,A., and Mura,U. (1998). Thioltransferase activity of bovine lens glutathione S-transferase. Biochem. J. 334 (Pt 1), 57-62.

287

Dalle-Donne,I., Milzani,A., Gagliano,N., Colombo,R., Giustarini,D., and Rossi,R. (2007). Molecular Mechanisms and Potential Clinical Significance of S- Glutathionylation. Antioxid. Redox. Signal.

Dan,H.C., Cooper,M.J., Cogswell,P.C., Duncan,J.A., Ting,J.P., and Baldwin,A.S. (2008). Akt-dependent regulation of NF-{kappa}B is controlled by mTOR and Raptor in association with IKK. Genes Dev. 22, 1490-1500.

Das,D.K. (2004). Thioredoxin regulation of ischemic preconditioning. Antioxid. Redox. Signal. 6, 405-412.

Davies,A.M., Malone,M.E., Martin,E.A., Jones,R.M., Jukes,R., Lim,C.K., Smith,L.L., and White,I.N. (1997). Peroxidase activation of 4-hydroxytamoxifen to free radicals detected by EPR spectroscopy. Free Radic. Biol. Med. 22, 423-431.

Davies,M.J. and Hawkins,C.L. (2000). Hypochlorite-induced oxidation of thiols: formation of thiyl radicals and the role of sulfenyl chlorides as intermediates. Free Radic. Res. 33, 719-729.

Davis,D.A., Newcomb,F.M., Starke,D.W., Ott,D.E., Mieyal,J.J., and Yarchoan,R. (1997). Thioltransferase (glutaredoxin) is detected within HIV-1 and can regulate the activity of glutathionylated HIV-1 protease in vitro. J. Biol. Chem. 272, 25935-25940.

Demasi,M., Silva,G.M., and Netto,L.E. (2003). 20 S proteasome from Saccharomyces cerevisiae is responsive to redox modifications and is S-glutathionylated. J. Biol. Chem. 278, 679-685.

Denecker,G., Ovaere,P., Vandenabeele,P., and Declercq,W. (2008). Caspase-14 reveals its secrets. J. Cell Biol. 180, 451-458.

Denu,J.M. and Tanner,K.G. (1998). Specific and reversible inactivation of protein tyrosine phosphatases by hydrogen peroxide: evidence for a sulfenic acid intermediate and implications for redox regulation. Biochemistry 37, 5633-5642.

Dhingra,S., Sharma,A.K., Arora,R.C., Slezak,J., and Singal,P.K. (2009). IL-10 attenuates TNF-alpha-induced NF kappaB pathway activation and cardiomyocyte apoptosis. Cardiovasc. Res. 82, 59-66.

Di Stefano,A., Frosali,S., Leonini,A., Ettorre,A., Priora,R., Di Simplicio,F.C., and Di Simplicio,P. (2006). GSH depletion, protein S-glutathionylation and mitochondrial transmembrane potential hyperpolarization are early events in initiation of cell death induced by a mixture of isothiazolinones in HL60 cells. Biochim. Biophys. Acta 1763, 214-225.

Discola,K.F., de Oliveira,M.A., Rosa,C., Jr., Monteiro,G., Barcena,J.A., Porras,P., Padilla,C.A., Guimaraes,B.G., and Soares Netto,L.E. (2008). Structural Aspects of the Distinct Biochemical Properties of Glutaredoxin 1 and Glutaredoxin 2 from Saccharomyces cerevisiae. J. Mol. Biol.

288

Dixon,W.G. and Symmons,D.P. (2007). What effects might anti-TNFalpha treatment be expected to have on cardiovascular morbidity and mortality in rheumatoid arthritis? A review of the role of TNFalpha in cardiovascular pathophysiology. Ann. Rheum. Dis. 66, 1132-1136.

Djurovic,S., Iversen,N., Jeansson,S., Hoover,F., and Christensen,G. (2004). Comparison of nonviral transfection and adeno-associated viral transduction on cardiomyocytes. Mol. Biotechnol. 28, 21-32.

Domenech,R.J., Macho,P., Velez,D., Sanchez,G., Liu,X., and Dhalla,N. (1998). Tachycardia preconditions infarct size in dogs: role of adenosine and protein kinase C. Circulation 97, 786-794.

Downey,J.M., Davis,A.M., and Cohen,M.V. (2007). Signaling pathways in ischemic preconditioning. Heart Fail. Rev 12, 181-188.

Duckert,P., Brunak,S., and Blom,N. (2004). Prediction of proprotein convertase cleavage sites. Protein Eng Des Sel 17, 107-112.

Eaton,P. and Shattock,M.J. (2002). Purification of proteins susceptible to oxidation at cysteine residues: identification of malate dehydrogenase as a target for S-glutathiolation. Ann. N. Y. Acad. Sci. 973, 529-532.

Eaton,P., Wright,N., Hearse,D.J., and Shattock,M.J. (2002). Glyceraldehyde phosphate dehydrogenase oxidation during cardiac ischemia and reperfusion. J. Mol. Cell Cardiol. 34, 1549-1560.

Eiserich,J.P., Patel,R.P., and O'Donnell,V.B. (1998). Pathophysiology of nitric oxide and related species: free radical reactions and modification of biomolecules. Mol. Aspects Med. 19, 221-357.

Elgan,T.H. and Berndt,K.D. (2008). Quantifying Escherichia coli Glutaredoxin-3 Substrate Specificity Using Ligand-induced Stability. J. Biol. Chem. 283, 32839-32847.

Enoksson,M., Fernandes,A.P., Prast,S., Lillig,C.H., Holmgren,A., and Orrenius,S. (2005). Overexpression of glutaredoxin 2 attenuates apoptosis by preventing cytochrome c release. Biochem. Biophys. Res. Commun. 327, 774-779.

Felder,C.E., Prilusky,J., Silman,I., and Sussman,J.L. (2007). A server and database for dipole moments of proteins. Nucleic Acids Res. 35, W512-W521.

Feng,Y., Zhong,N., Rouhier,N., Hase,T., Kusunoki,M., Jacquot,J.P., Jin,C., and Xia,B. (2006). Structural insight into poplar glutaredoxin C1 with a bridging iron-sulfur cluster at the active site. Biochemistry 45, 7998-8008.

Fernandes,A.P., Fladvad,M., Berndt,C., Andresen,C., Lillig,C.H., Neubauer,P., Sunnerhagen,M., Holmgren,A., and Vlamis-Gardikas,A. (2005). A novel monothiol

289

glutaredoxin (Grx4) from Escherichia coli can serve as a substrate for thioredoxin reductase. J. Biol. Chem. 280, 24544-24552.

Fernandes,A.P. and Holmgren,A. (2004). Glutaredoxins: glutathione-dependent redox enzymes with functions far beyond a simple thioredoxin backup system. Antioxid. Redox. Signal. 6, 63-74.

Fernando,M.R., Lechner,J.M., Lofgren,S., Gladyshev,V.N., and Lou,M.F. (2006). Mitochondrial thioltransferase (glutaredoxin 2) has GSH-dependent and thioredoxin reductase-dependent peroxidase activities in vitro and in lens epithelial cells. FASEB J. 20, 2645-2647.

Fiebig,A.A., Zhu,W., Hollerbach,C., Leber,B., and Andrews,D.W. (2006). Bcl-XL is qualitatively different from and ten times more effective than Bcl-2 when expressed in a breast cancer cell line. BMC. Cancer 6, 213.

Figueiredo-Pereira,M.E., Yakushin,S., and Cohen,G. (1998). Disruption of the intracellular sulfhydryl homeostasis by cadmium-induced oxidative stress leads to protein thiolation and ubiquitination in neuronal cells. J. Biol. Chem. 273, 12703-12709.

Findlay,V.J., Townsend,D.M., Morris,T.E., Fraser,J.P., He,L., and Tew,K.D. (2006). A novel role for human sulfiredoxin in the reversal of glutathionylation. Cancer Res. 66, 6800-6806.

Findlay,V.J., Townsend,D.M., Saavedra,J.E., Buzard,G.S., Citro,M.L., Keefer,L.K., Ji,X., and Tew,K.D. (2004). Tumor cell responses to a novel glutathione S-transferase- activated nitric oxide-releasing prodrug. Mol. Pharmacol. 65, 1070-1079.

Finkel,T. (2000). Redox-dependent signal transduction. FEBS Lett. 476, 52-54.

Fladvad,M., Bellanda,M., Fernandes,A.P., Mammi,S., Vlamis-Gardikas,A., Holmgren,A., and Sunnerhagen,M. (2005). Molecular mapping of functionalities in the solution structure of reduced Grx4, a monothiol glutaredoxin from Escherichia coli. J. Biol. Chem. 280, 24553-24561.

Flohe,L. (1985). The glutathione peroxidase reaction: molecular basis of the antioxidant function of selenium in mammals. Curr. Top. Cell Regul. 27, 473-478.

Flotho,C., Coustan-Smith,E., Pei,D., Iwamoto,S., Song,G., Cheng,C., Pui,C.H., Downing,J.R., and Campana,D. (2006). Genes contributing to minimal residual disease in childhood acute lymphoblastic leukemia: prognostic significance of CASP8AP2. Blood 108, 1050-1057.

Foloppe,N. and Nilsson,L. (2007). Stabilization of the catalytic thiolate in a mammalian glutaredoxin: structure, dynamics and electrostatics of reduced pig glutaredoxin and its mutants. J. Mol. Biol. 372, 798-816.

290

Frolkis,V.V., Frolkis,R.A., Mkhitarian,L.S., and Fraifeld,V.E. (1991). Age-dependent effects of ischemia and reperfusion on cardiac function and Ca2+ transport in myocardium. Gerontology 37, 233-239.

Gallogly,M.M. and Mieyal,J.J. (2007). Mechanisms of reversible protein glutathionylation in redox signaling and oxidative stress. Curr. Opin. Pharmacol. 7, 381- 391.

Gallogly,M.M., Starke,D.W., Leonberg,A.K., Ospina,S.M., and Mieyal,J.J. (2008). Kinetic and mechanistic characterization and versatile catalytic properties of mammalian glutaredoxin 2: implications for intracellular roles. Biochemistry 47, 11144-11157.

Gallogly,M.M., Starke,D.W., and Mieyal,J.J. (2009). Mechanistic and Kinetic Details of Thiol-Disulfide Exchange by Glutaredoxins and Potential Mechanisms of Regulation. Antioxid. Redox. Signal.

Gan,Z.R. (1992). Cloning and sequencing of a gene encoding yeast thioltransferase. Biochem. Biophys. Res. Commun. 187, 949-955.

Gan,Z.R. and Wells,W.W. (1986). Purification and properties of thioltransferase. J. Biol. Chem. 261, 996-1001.

Gan,Z.R. and Wells,W.W. (1987). Identification and reactivity of the catalytic site of pig liver thioltransferase. J. Biol. Chem. 262, 6704-6707.

Gao,Z.Q., Yang,C., Wang,Y.Y., Wang,P., Chen,H.L., Zhang,X.D., Liu,R., Li,W.L., Qin,X.J., Liang,X., and Hai,C.X. (2008). RAGE upregulation and nuclear factor-kappaB activation associated with ageing rat cardiomyocyte dysfunction. Gen. Physiol Biophys. 27, 152-158.

Garcera,A., Barreto,L., Piedrafita,L., Tamarit,J., and Herrero,E. (2006). Saccharomyces cerevisiae cells have three Omega class glutathione S-transferases acting as 1-Cys thiol transferases. Biochem. J. 398, 187-196.

Gaston,B. (1999). Nitric oxide and thiol groups. Biochim. Biophys. Acta 1411, 323-333.

Ghezzi,P. (2005a). Oxidoreduction of protein thiols in redox regulation. Biochem. Soc. Trans. 33, 1378-1381.

Ghezzi,P. (2005b). Regulation of protein function by glutathionylation. Free Radic. Res. 39, 573-580.

Gilbert,H.F. (1990). Molecular and cellular aspects of thiol-disulfide exchange. Adv. Enzymol. Relat Areas Mol. Biol. 63, 69-172.

Gilbert,H.F. (1995). Thiol/disulfide exchange equilibria and disulfide bond stability. Methods Enzymol. 251, 8-28.

291

Giustarini,D., Milzani,A., Aldini,G., Carini,M., Rossi,R., and Dalle-Donne,I. (2005). S- nitrosation versus S-glutathionylation of protein sulfhydryl groups by S- nitrosoglutathione. Antioxid. Redox. Signal. 7, 930-939.

Giustarini,D., Rossi,R., Milzani,A., Colombo,R., and Dalle-Donne,I. (2004). S- glutathionylation: from redox regulation of protein functions to human diseases. J. Cell Mol. Med. 8, 201-212.

Gladyshev,V.N., Liu,A., Novoselov,S.V., Krysan,K., Sun,Q.A., Kryukov,V.M., Kryukov,G.V., and Lou,M.F. (2001). Identification and characterization of a new mammalian glutaredoxin (thioltransferase), Grx2. J. Biol. Chem. 276, 30374-30380.

Gomes,D.S., Pereira,M.D., Panek,A.D., Andrade,L.R., and Eleutherio,E.C. (2008). Apoptosis as a mechanism for removal of mutated cells of Saccharomyces cerevisiae: the role of Grx2 under cadmium exposure. Biochim. Biophys. Acta 1780, 160-166.

Gravina,S.A. and Mieyal,J.J. (1993). Thioltransferase is a specific glutathionyl mixed disulfide oxidoreductase. Biochemistry 32, 3368-3376.

Gross,A., McDonnell,J.M., and Korsmeyer,S.J. (1999). BCL-2 family members and the mitochondria in apoptosis. Genes Dev. 13, 1899-1911.

Gustafsson,A.B. and Gottlieb,R.A. (2007). Bcl-2 family members and apoptosis, taken to heart. Am. J. Physiol Cell Physiol 292, C45-C51.

Hakansson,K.O. and Winther,J.R. (2007). Structure of glutaredoxin Grx1p C30S mutant from yeast. Acta Crystallogr. D. Biol. Crystallogr. 63, 288-294.

Halestrap,A.P., Woodfield,K.Y., and Connern,C.P. (1997). Oxidative stress, thiol reagents, and membrane potential modulate the mitochondrial permeability transition by affecting nucleotide binding to the adenine nucleotide translocase. J. Biol. Chem. 272, 3346-3354.

Halliwell,B. (1996). Mechanisms involved in the generation of free radicals. Pathol. Biol. (Paris) 44, 6-13.

Han,D., Canali,R., Garcia,J., Aguilera,R., Gallaher,T.K., and Cadenas,E. (2005). Sites and mechanisms of aconitase inactivation by peroxynitrite: modulation by citrate and glutathione. Biochemistry 44, 11986-11996.

Han,H., Long,H., Wang,H., Wang,J., Zhang,Y., and Wang,Z. (2004). Progressive apoptotic cell death triggered by transient oxidative insult in H9c2 rat ventricular cells: a novel pattern of apoptosis and the mechanisms. Am. J. Physiol Heart Circ. Physiol 286, H2169-H2182.

Harman,L.S., Carver,D.K., Schreiber,J., and Mason,R.P. (1986). One- and two-electron oxidation of reduced glutathione by peroxidases. J. Biol. Chem. 261, 1642-1648.

292

Harrison,D., gongora,M.C., Guzik,T.J., and Widder,J. (2007). Oxidative Stress and Hypertension. J Am Soc Hypert 1, 30-44.

Hashemy,S.I., Johansson,C., Berndt,C., Lillig,C.H., and Holmgren,A. (2007). Oxidation and S-nitrosylation of cysteines in human cytosolic and mitochondrial glutaredoxins: effects on structure and activity. J. Biol. Chem. 282, 14428-14436.

Hatakeyama,M., Tanimoto,Y., and Mizoguchi,T. (1984). Purification and some properties of bovine liver cytosol thioltransferase. J. Biochem. 95, 1811-1818.

Hausenloy,D.J. and Yellon,D.M. (2006). Survival kinases in ischemic preconditioning and postconditioning. Cardiovasc. Res. 70, 240-253.

Hayes,J.D., Flanagan,J.U., and Jowsey,I.R. (2005). Glutathione transferases. Annu. Rev. Pharmacol. Toxicol. 45, 51-88.

Heineke,J. and Molkentin,J.D. (2006). Regulation of cardiac hypertrophy by intracellular signalling pathways. Nat. Rev. Mol. Cell Biol. 7, 589-600.

Hewitt,S.M., Hamada,S., McDonnell,T.J., Rauscher,F.J., III, and Saunders,G.F. (1995). Regulation of the proto-oncogenes bcl-2 and c-myc by the Wilms' tumor suppressor gene WT1. Cancer Res. 55, 5386-5389.

Hirota,K., Matsui,M., Murata,M., Takashima,Y., Cheng,F.S., Itoh,T., Fukuda,K., and Yodoi,J. (2000). Nucleoredoxin, glutaredoxin, and thioredoxin differentially regulate NF- kappaB, AP-1, and CREB activation in HEK293 cells. Biochem. Biophys. Res. Commun. 274, 177-182.

Hirst,J., Carroll,J., Fearnley,I.M., Shannon,R.J., and Walker,J.E. (2003). The nuclear encoded subunits of complex I from bovine heart mitochondria. Biochim. Biophys. Acta 1604, 135-150.

Ho,Y.S., Xiong,Y., Ho,D.S., Gao,J., Chua,B.H., Pai,H., and Mieyal,J.J. (2007). Targeted disruption of the glutaredoxin 1 gene does not sensitize adult mice to tissue injury induced by ischemia/reperfusion and hyperoxia. Free Radic. Biol. Med. 43, 1299-1312.

Hoffman,B. and Liebermann,D.A. (2009). Gadd45 modulation of intrinsic and extrinsic stress responses in myeloid cells. J. Cell Physiol 218, 26-31.

Hogg, D. R. Sulphenic Acids and Their Derivatives. Jones, D. N. 262-310. 1979. Pergamon Press. Comprehensive Organic Chemistry: The Synthesis and Reactions of Organic Compounds. Ref Type: Serial (Book,Monograph)

Holmgren,A. (1976). Hydrogen donor system for Escherichia coli ribonucleoside- diphosphate reductase dependent upon glutathione. Proc. Natl. Acad. Sci. U. S. A 73, 2275-2279.

293

Holmgren,A. (1985). Thioredoxin. Annu. Rev. Biochem. 54, 237-271.

Hotchkiss,R.S., McConnell,K.W., Bullock,K.B., Davis,C.G., Chang,K.C., Schwulst,S.J., Dunne J.C., Dietz,G.P.H., Bähr,M., McDunn,J.E., Karl,I.E., Wagner,T.H., Perren Cobb,J., Coopersmith,C.M., Piwnica-Worms,D. (2006). TAT-BH4 and TAT-Bcl-xL Peptides Protect against Sepsis-Induced Lymphocyte Apoptosis In Vivo. J. Immunol. 176, 5471-5477.

Hsu,M., Srinivas,B., Kumar,J., Subramanian,R., and Andersen,J. (2005). Glutathione depletion resulting in selective mitochondrial complex I inhibition in dopaminergic cells is via an NO-mediated pathway not involving peroxynitrite: implications for Parkinson's disease. J. Neurochem. 92, 1091-1103.

Hu,Y., Benedict,M.A., Wu,D., Inohara,N., and Nunez,G. (1998). Bcl-XL interacts with Apaf-1 and inhibits Apaf-1-dependent caspase-9 activation. Proc. Natl. Acad. Sci. U. S. A 95, 4386-4391.

Huang,J., Ito,Y., Morikawa,M., Uchida,H., Kobune,M., Sasaki,K., Abe,T., and Hamada,H. (2003). Bcl-xL gene transfer protects the heart against ischemia/reperfusion injury. Biochem. Biophys. Res. Commun. 311, 64-70.

Huang,K.P. and Huang,F.L. (2002). Glutathionylation of proteins by glutathione disulfide S-oxide. Biochem. Pharmacol. 64, 1049-1056.

Hudemann,C., Lonn,M.E., Godoy,J.R., Avval,F.Z., Capani,F., Holmgren,A., and Lillig,C.H. (2009). Identification, expression pattern, and characterization of mouse glutaredoxin 2 isoforms. Antioxid. Redox. Signal. 11, 1-14.

Humphries,K.M., Juliano,C., and Taylor,S.S. (2002). Regulation of cAMP-dependent protein kinase activity by glutathionylation. J Biol. Chem. 277, 43505-43511.

Hurd,T.R., Costa,N.J., Dahm,C.C., Beer,S.M., Brown,S.E., Filipovska,A., and Murphy,M.P. (2005). Glutathionylation of mitochondrial proteins. Antioxid. Redox. Signal. 7, 999-1010.

Hurd,T.R., Requejo,R., Filipovska,A., Brown,S., Prime,T.A., Robinson,A.J., Fearnley,I.M., and Murphy,M.P. (2008). Complex I within oxidatively stressed bovine heart mitochondria is glutathionylated on Cys-531 and Cys-704 of the 75-kDa subunit: potential role of CYS residues in decreasing oxidative damage. J. Biol. Chem. 283, 24801-24815.

Imahashi,K., Schneider,M.D., Steenbergen,C., and Murphy,E. (2004). Transgenic expression of Bcl-2 modulates energy metabolism, prevents cytosolic acidification during ischemia, and reduces ischemia/reperfusion injury. Circ. Res. 95, 734-741.

Imlay,J.A. (2006). Iron-sulphur clusters and the problem with oxygen. Mol. Microbiol. 59, 1073-1082.

294

Isakov,N., Witte,S., and Altman,A. (2000). PICOT-HD: a highly conserved protein domain that is often associated with thioredoxin and glutaredoxin modules. Trends Biochem. Sci. 25, 537-539.

Ishitani,R., Tajima,H., Takata,H., Tsuchiya,K., Kuwae,T., Yamada,M., Takahashi,H., Tatton,N.A., and Katsube,N. (2003). Proapoptotic protein glyceraldehyde-3-phosphate dehydrogenase: a possible site of action of antiapoptotic drugs. Prog. Neuropsychopharmacol. Biol. Psychiatry 27, 291-301.

Isoyama,S., Wei,J.Y., Izumo,S., Fort,P., Schoen,F.J., and Grossman,W. (1987). Effect of age on the development of cardiac hypertrophy produced by aortic constriction in the rat. Circ. Res. 61, 337-345.

Jaeschke,A., Czech,M.P., and Davis,R.J. (2004). An essential role of the JIP1 scaffold protein for JNK activation in adipose tissue. Genes Dev. 18, 1976-1980.

Jahngen-Hodge,J., Obin,M.S., Gong,X., Shang,F., Nowell,T.R., Jr., Gong,J., Abasi,H., Blumberg,J., and Taylor,A. (1997). Regulation of ubiquitin-conjugating enzymes by glutathione following oxidative stress. J. Biol. Chem. 272, 28218-28226.

Jao,S.C., English Ospina,S.M., Berdis,A.J., Starke,D.W., Post,C.B., and Mieyal,J.J. (2006). Computational and mutational analysis of human glutaredoxin (thioltransferase): probing the molecular basis of the low pKa of cysteine 22 and its role in catalysis. Biochemistry 45, 4785-4796.

Jeong,W., Park,S.J., Chang,T.S., Lee,D.Y., and Rhee,S.G. (2006). Molecular mechanism of the reduction of cysteine sulfinic acid of peroxiredoxin to cysteine by mammalian sulfiredoxin. J. Biol. Chem. 281, 14400-14407.

Ji,Y., Akerboom,T.P., Sies,H., and Thomas,J.A. (1999). S-nitrosylation and S- glutathiolation of protein sulfhydryls by S-nitroso glutathione. Arch. Biochem. Biophys. 362, 67-78.

Johansson,C., Kavanagh,K.L., Gileadi,O., and Oppermann,U. (2007). Reversible sequestration of active site cysteines in a 2Fe-2S-bridged dimer provides a mechanism for glutaredoxin 2 regulation in human mitochondria. J. Biol. Chem. 282, 3077-3082.

Johansson,C., Lillig,C.H., and Holmgren,A. (2004). Human mitochondrial glutaredoxin reduces S-glutathionylated proteins with high affinity accepting electrons from either glutathione or thioredoxin reductase. J. Biol. Chem. 279, 7537-7543.

Jones,W.K., Brown,M., Ren,X., He,S., and McGuinness,M. (2003). NF-kappaB as an integrator of diverse signaling pathways: the heart of myocardial signaling? Cardiovasc. Toxicol. 3, 229-254.

Jones,W.K., Brown,M., Wilhide,M., He,S., and Ren,X. (2005). NF-kappaB in cardiovascular disease: diverse and specific effects of a "general" transcription factor? Cardiovasc. Toxicol. 5, 183-202.

295

Juhaszova,M., Rabuel,C., Zorov,D.B., Lakatta,E.G., and Sollott,S.J. (2005). Protection in the aged heart: preventing the heart-break of old age? Cardiovasc. Res. 66, 233-244.

Jung,C.H. and Thomas,J.A. (1996). S-glutathiolated hepatocyte proteins and insulin disulfides as substrates for reduction by glutaredoxin, thioredoxin, protein disulfide isomerase, and glutathione. Arch. Biochem. Biophys. 335, 61-72.

Kajstura,J., Cheng,W., Sarangarajan,R., Li,P., Li,B., Nitahara,J.A., Chapnick,S., Reiss,K., Olivetti,G., and Anversa,P. (1996). Necrotic and apoptotic myocyte cell death in the aging heart of Fischer 344 rats. Am. J. Physiol 271, H1215-H1228.

Kallis,G.B. and Holmgren,A. (1980). Differential reactivity of the functional sulfhydryl groups of cysteine-32 and cysteine-35 present in the reduced form of thioredoxin from Escherichia coli. J. Biol. Chem. 255, 10261-10265.

Kalogeropoulos,A., Georgiopoulou,V., Kritchevsky,S.B., Psaty,B.M., Smith,N.L., Newman,A.B., Rodondi,N., Satterfield,S., Bauer,D.C., Bibbins-Domingo,K., Smith,A.L., Wilson,P.W., Vasan,R.S., Harris,T.B., and Butler,J. (2009). Epidemiology of incident heart failure in a contemporary elderly cohort: the health, aging, and body composition study. Arch. Intern. Med. 169, 708-715.

Kanda,M., Ihara,Y., Murata,H., Urata,Y., Kono,T., Yodoi,J., Seto,S., Yano,K., and Kondo,T. (2006). Glutaredoxin modulates platelet-derived growth factor-dependent cell signaling by regulating the redox status of low molecular weight protein-tyrosine phosphatase. J. Biol. Chem. 281, 28518-28528.

Kane,D.J., Sarafian,T.A., Anton,R., Hahn,H., Gralla,E.B., Valentine,J.S., Ord,T., and Bredesen,D.E. (1993). Bcl-2 inhibition of neural death: decreased generation of reactive oxygen species. Science 262, 1274-1277.

Karoui,H., Hogg,N., Frejaville,C., Tordo,P., and Kalyanaraman,B. (1996). Characterization of sulfur-centered radical intermediates formed during the oxidation of thiols and sulfite by peroxynitrite. ESR-spin trapping and oxygen uptake studies. J. Biol. Chem. 271, 6000-6009.

Kenchappa,R.S., Diwakar,L., Annepu,J., and Ravindranath,V. (2004). Estrogen and neuroprotection: higher constitutive expression of glutaredoxin in female mice offers protection against MPTP-mediated neurodegeneration. FASEB J. 18, 1102-1104.

Kher,N. and Marsh,J.D. (2004). Pathobiology of atherosclerosis--a brief review. Semin. Thromb. Hemost. 30, 665-672.

Kim,J.S., Jin,Y., and Lemasters,J.J. (2006). Reactive oxygen species, but not Ca2+ overloading, trigger pH- and mitochondrial permeability transition-dependent death of adult rat myocytes after ischemia-reperfusion. Am. J. Physiol Heart Circ. Physiol 290, H2024-H2034.

296

Kim,R. (2005). Unknotting the roles of Bcl-2 and Bcl-xL in cell death. Biochem. Biophys. Res. Commun. 333, 336-343.

Kim,S.J., Chung,Y.K., Chung,T.W., Kim,J.R., Moon,S.K., Kim,C.H., and Park,Y.G. (2009). Regulation of matrix metalloproteinase-9 expression between gingival fibroblast cells from old and young rats. Biochem. Biophys. Res. Commun. 378, 152-156.

Klatt,P. and Lamas,S. (2000). Regulation of protein function by S-glutathiolation in response to oxidative and nitrosative stress. Eur. J. Biochem. 267, 4928-4944.

Klatt,P., Molina,E.P., De Lacoba,M.G., Padilla,C.A., Martinez-Galesteo,E., Barcena,J.A., and Lamas,S. (1999a). Redox regulation of c-Jun DNA binding by reversible S- glutathiolation. FASEB J. 13, 1481-1490.

Klatt,P., Molina,E.P., and Lamas,S. (1999b). Nitric oxide inhibits c-Jun DNA binding by specifically targeted S-glutathionylation. J. Biol. Chem. 274, 15857-15864.

Kobayashi,S., Lackey,T., Huang,Y., Bisping,E., Pu,W.T., Boxer,L.M., and Liang,Q. (2006). Transcription factor gata4 regulates cardiac BCL2 gene expression in vitro and in vivo. FASEB J. 20, 800-802.

Kortemme,T. and Creighton,T.E. (1995). Ionisation of cysteine residues at the termini of model alpha-helical peptides. Relevance to unusual thiol pKa values in proteins of the thioredoxin family. J. Mol. Biol. 253, 799-812.

Koul,D., Yao,Y., Abbruzzese,J.L., Yung,W.K., and Reddy,S.A. (2001). Tumor suppressor MMAC/PTEN inhibits cytokine-induced NFkappaB activation without interfering with the IkappaB degradation pathway. J. Biol. Chem. 276, 11402-11408.

Krebs,J.F., Armstrong,R.C., Srinivasan,A., Aja,T., Wong,A.M., Aboy,A., Sayers,R., Pham,B., Vu,T., Hoang,K., Karanewsky,D.S., Leist,C., Schmitz,A., Wu,J.C., Tomaselli,K.J., and Fritz,L.C. (1999). Activation of membrane-associated procaspase-3 is regulated by Bcl-2. J. Cell Biol. 144, 915-926.

Kuo,Y.Y. and Chang,Z.F. (2007). GATA-1 and Gfi-1B interplay to regulate Bcl-xL transcription. Mol. Cell Biol. 27, 4261-4272.

Kwak,H.B., Song,W., and Lawler,J.M. (2006). Exercise training attenuates age-induced elevation in Bax/Bcl-2 ratio, apoptosis, and remodeling in the rat heart. FASEB J. 20, 791-793.

Kwak,H.S., Yim,H.S., Chock,P.B., and Yim,M.B. (1995). Endogenous intracellular glutathionyl radicals are generated in neuroblastoma cells under hydrogen peroxide oxidative stress. Proc. Natl. Acad. Sci. U. S. A 92, 4582-4586.

Landmesser,U., Cai,H., Dikalov,S., McCann,L., Hwang,J., Jo,H., Holland,S.M., and Harrison,D.G. (2002). Role of p47(phox) in vascular oxidative stress and hypertension caused by angiotensin II. Hypertension 40, 511-515.

297

Lazou,A., Iliodromitis,E.K., Cieslak,D., Voskarides,K., Mousikos,S., Bofilis,E., and Kremastinos,D.T. (2006). Ischemic but not mechanical preconditioning attenuates ischemia/reperfusion induced myocardial apoptosis in anaesthetized rabbits: the role of Bcl-2 family proteins and ERK1/2. Apoptosis. 11, 2195-2204.

Lee,R.T. and Libby,P. (1997). The unstable atheroma. Arterioscler. Thromb. Vasc. Biol. 17, 1859-1867.

Lee,S.R., Kwon,K.S., Kim,S.R., and Rhee,S.G. (1998). Reversible inactivation of protein-tyrosine phosphatase 1B in A431 cells stimulated with epidermal growth factor. J. Biol. Chem. 273, 15366-15372.

Lee,Y. and Gustafsson,A.B. (2009). Role of apoptosis in cardiovascular disease. Apoptosis. 14, 536-548.

Lees,S.J., Zwetsloot,K.A., and Booth,F.W. (2009). Muscle precursor cells isolated from aged rats exhibit an increased tumor necrosis factor- alpha response. Aging Cell 8, 26-35.

Lesnefsky,E.J., Chen,Q., Moghaddas,S., Hassan,M.O., Tandler,B., and Hoppel,C.L. (2004). Blockade of electron transport during ischemia protects cardiac mitochondria. J Biol. Chem. 279, 47961-47967.

Lesnefsky,E.J., Gallo,D.S., Ye,J., Whittingham,T.S., and Lust,W.D. (1994). Aging increases ischemia-reperfusion injury in the isolated, buffer-perfused heart. J. Lab Clin. Med. 124, 843-851.

Lesnefsky,E.J. and Hoppel,C.L. (2003). Ischemia-reperfusion injury in the aged heart: role of mitochondria. Arch. Biochem. Biophys. 420, 287-297.

Lesnefsky,E.J., Lundergan,C.F., Hodgson,J.M., Nair,R., Reiner,J.S., Greenhouse,S.W., Califf,R.M., and Ross,A.M. (1996). Increased left ventricular dysfunction in elderly patients despite successful thrombolysis: the GUSTO-I angiographic experience. J. Am. Coll. Cardiol. 28, 331-337.

Li,J., Huang,F.L., and Huang,K.P. (2001). Glutathiolation of proteins by glutathione disulfide S-oxide derived from S-nitrosoglutathione. Modifications of rat brain neurogranin/RC3 and neuromodulin/GAP-43. J. Biol. Chem. 276, 3098-3105.

Li,Z.L., Abe,H., Ueki,K., Kumagai,K., Araki,R., and Otsuki,Y. (2003). Identification of c-Jun as bcl-2 transcription factor in human uterine endometrium. J. Histochem. Cytochem. 51, 1601-1609.

Lieber,S.C., Aubry,N., Pain,J., Diaz,G., Kim,S.J., and Vatner,S.F. (2004). Aging increases stiffness of cardiac myocytes measured by atomic force microscopy nanoindentation. Am. J. Physiol Heart Circ. Physiol 287, H645-H651.

Lillig,C.H., Berndt,C., and Holmgren,A. (2008). Glutaredoxin systems. Biochim. Biophys. Acta 1780, 1304-1317.

298

Lillig,C.H., Berndt,C., Vergnolle,O., Lonn,M.E., Hudemann,C., Bill,E., and Holmgren,A. (2005). Characterization of human glutaredoxin 2 as iron-sulfur protein: a possible role as redox sensor. Proc. Natl. Acad. Sci. U. S. A 102, 8168-8173.

Lillig,C.H., Lonn,M.E., Enoksson,M., Fernandes,A.P., and Holmgren,A. (2004). Short interfering RNA-mediated silencing of glutaredoxin 2 increases the sensitivity of HeLa cells toward doxorubicin and phenylarsine oxide. Proc. Natl. Acad. Sci. U. S. A 101, 13227-13232.

Lind,C., Gerdes,R., Hamnell,Y., Schuppe-Koistinen,I., von Lowenhielm,H.B., Holmgren,A., and Cotgreave,I.A. (2002). Identification of S-glutathionylated cellular proteins during oxidative stress and constitutive metabolism by affinity purification and proteomic analysis. Arch. Biochem. Biophys. 406, 229-240.

Lipman,D.J. and Pearson,W.R. (1985). Rapid and sensitive protein similarity searches. Science. 227, 1435-1441.

Liu,P., Xu,B., Cavalieri,T.A., and Hock,C.E. (2002). Age-related difference in myocardial function and inflammation in a rat model of myocardial ischemia-reperfusion. Cardiovasc. Res. 56, 443-453.

Livak,K.J., Schmittgen,T.D. (2001). Analysis of relative gene expression data using real- −ΔΔC time quantitative PCR and the 2 T method. Methods 25, 402-408.

Llopis,J., McCaffery,J.M., Miyawaki,A., Farquhar,M.G., and Tsien,R.Y. (1998). Measurement of cytosolic, mitochondrial, and Golgi pH in single living cells with green fluorescent proteins. Proc. Natl. Acad. Sci. U. S. A 95, 6803-6808.

Lobysheva,I.I., Serezhenkov,V.A., and Vanin,A.F. (1999). Interaction of peroxynitrite and hydrogen peroxide with dinitrosyl iron complexes containing thiol ligands in vitro. Biochemistry (Mosc. ) 64, 153-158.

Lonn,M.E., Hudemann,C., Berndt,C., Cherkasov,V., Capani,F., Holmgren,A., and Lillig,C.H. (2007). Expression Pattern of Human Glutaredoxin 2 Isoforms: Identification and Characterization of Two Testis/Cancer Cell-Specific Isoforms. Antioxid. Redox. Signal.

Lonn,M.E., Hudemann,C., Berndt,C., Cherkasov,V., Capani,F., Holmgren,A., and Lillig,C.H. (2008). Expression pattern of human glutaredoxin 2 isoforms: identification and characterization of two testis/cancer cell-specific isoforms. Antioxid. Redox. Signal. 10, 547-557.

Luikenhuis,S., Perrone,G., Dawes,I.W., and Grant,C.M. (1998). The yeast Saccharomyces cerevisiae contains two glutaredoxin genes that are required for protection against reactive oxygen species. Mol. Biol. Cell 9, 1081-1091.

299

Lundberg,M., Fernandes,A.P., Kumar,S., and Holmgren,A. (2004). Cellular and plasma levels of human glutaredoxin 1 and 2 detected by sensitive ELISA systems. Biochem. Biophys. Res. Commun. 319, 801-809.

Lundberg,M., Holmgren,A., and Johansson,M. (2006). Human glutaredoxin 2 affinity tag for recombinant peptide and protein purification. Protein Expr. Purif. 45, 37-42.

Lundberg,M., Johansson,C., Chandra,J., Enoksson,M., Jacobsson,G., Ljung,J., Johansson,M., and Holmgren,A. (2001). Cloning and expression of a novel human glutaredoxin (Grx2) with mitochondrial and nuclear isoforms. J. Biol. Chem. 276, 26269- 26275.

Luthman,M. and Holmgren,A. (1982). Glutaredoxin from calf thymus. Purification to homogeneity. J. Biol. Chem. 257, 6686-6690.

Malik,G., Nagy,N., Ho,Y.S., Maulik,N., and Das,D.K. (2008). Role of glutaredoxin-1 in cardioprotection: an insight with Glrx1 transgenic and knockout animals. J. Mol. Cell Cardiol. 44, 261-269.

Manevich,Y., Feinstein,S.I., and Fisher,A.B. (2004). Activation of the antioxidant enzyme 1-CYS peroxiredoxin requires glutathionylation mediated by heterodimerization with pi GST. Proc. Natl. Acad. Sci. U. S. A 101, 3780-3785.

Mani,K. (2008). Programmed cell death in cardiac myocytes: strategies to maximize post-ischemic salvage. Heart Fail. Rev. 13, 193-209.

Maples,K.R., Kennedy,C.H., Jordan,S.J., and Mason,R.P. (1990). In vivo thiyl free radical formation from hemoglobin following administration of hydroperoxides. Arch. Biochem. Biophys. 277, 402-409.

Martinez-Vicente,M., Sovak,G., and Cuervo,A.M. (2005). Protein degradation and aging. Exp. Gerontol. 40, 622-633.

Maulik,N., Engelman,R.M., Rousou,J.A., Flack,J.E., III, Deaton,D., and Das,D.K. (1999). Ischemic preconditioning reduces apoptosis by upregulating anti-death gene Bcl- 2. Circulation 100, II369-II375.

Maulik,N., Sato,M., Price,B.D., and Das,D.K. (1998). An essential role of NFkappaB in tyrosine kinase signaling of p38 MAP kinase regulation of myocardial adaptation to ischemia. FEBS Lett. 429, 365-369.

Mayo,M.W., Madrid,L.V., Westerheide,S.D., Jones,D.R., Yuan,X.J., Baldwin,A.S., Jr., and Whang,Y.E. (2002). PTEN blocks tumor necrosis factor-induced NF-kappa B- dependent transcription by inhibiting the transactivation potential of the p65 subunit. J. Biol. Chem. 277, 11116-11125.

Mesecke,N., Mittler,S., Eckers,E., Herrmann,J.M., and Deponte,M. (2008). Two novel monothiol glutaredoxins from Saccharomyces cerevisiae provide further insight into iron-

300

sulfur cluster binding, oligomerization, and enzymatic activity of glutaredoxins. Biochemistry 47, 1452-1463.

Mieyal,J.J., Gallogly,M.M., Qanungo,S., Sabens,E.A., and Shelton,M.D. (2008). Molecular mechanisms and clinical implications of reversible protein S-glutathionylation. Antioxid. Redox. Signal. 10, 1941-1988.

Mieyal, J. J., Srinivasan, U., Starke, D. W., Gravina, S. A., and Mieyal, P. A. Glutathionyl specificity of the Thioltransferases: Mechanistic and Physiological Implications. Packer, L and Cadenas, E. 305-372. 1995. Marcel Dekker, Inc. Biothiols in Health and Disease. Ref Type: Serial (Book,Monograph)

Mieyal,J.J., Starke,D.W., Gravina,S.A., Dothey,C., and Chung,J.S. (1991a). Thioltransferase in human red blood cells: purification and properties. Biochemistry 30, 6088-6097.

Mieyal,J.J., Starke,D.W., Gravina,S.A., and Hocevar,B.A. (1991b). Thioltransferase in human red blood cells: kinetics and equilibrium. Biochemistry 30, 8883-8891.

Miyamoto,S., Murphy,A.N., and Brown,J.H. (2009). Akt mediated mitochondrial protection in the heart: metabolic and survival pathways to the rescue. J. Bioenerg. Biomembr.

Mohr,S., Hallak,H., de Boitte,A., Lapetina,E.G., and Brune,B. (1999). Nitric oxide- induced S-glutathionylation and inactivation of glyceraldehyde-3-phosphate dehydrogenase. J. Biol. Chem. 274, 9427-9430.

Molina-Navarro,M.M., Casas,C., Piedrafita,L., Belli,G., and Herrero,E. (2006). Prokaryotic and eukaryotic monothiol glutaredoxins are able to perform the functions of Grx5 in the biogenesis of Fe/S clusters in yeast mitochondria. FEBS Lett. 580, 2273- 2280.

Moore,K.W., de Waal,M.R., Coffman,R.L., and O'Garra,A. (2001). Interleukin-10 and the interleukin-10 receptor. Annu. Rev. Immunol. 19, 683-765.

Moorjani,N., Catarino,P., Trabzuni,D., Saleh,S., Moorji,A., Dzimiri,N., Al-Mohanna,F., Westaby,S., and Ahmad,M. (2007). Upregulation of Bcl-2 proteins during the transition to pressure overload-induced heart failure. Int. J. Cardiol. 116, 27-33.

Morell,S., Follmann,H., and Haberlein,I. (1995). Identification and localization of the first glutaredoxin in leaves of a higher plant. FEBS Lett. 369, 149-152.

Muhlenhoff,U., Gerber,J., Richhardt,N., and Lill,R. (2003). Components involved in assembly and dislocation of iron-sulfur clusters on the scaffold protein Isu1p. EMBO J. 22, 4815-4825.

301

Mukherjee,S., Gangopadhyay,H., and Das,D.K. (2008). Broccoli: a unique vegetable that protects mammalian hearts through the redox cycling of the thioredoxin superfamily. J. Agric. Food Chem. 56, 609-617.

Mullonkal,C.J. and Toledo-Pereyra,L.H. (2007). Akt in ischemia and reperfusion. J. Invest Surg. 20, 195-203.

Murata,H., Ihara,Y., Nakamura,H., Yodoi,J., Sumikawa,K., and Kondo,T. (2003). Glutaredoxin exerts an antiapoptotic effect by regulating the redox state of Akt. J. Biol. Chem. 278, 50226-50233.

Murry,C.E., Jennings,R.B., and Reimer,K.A. (1986). Preconditioning with ischemia: a delay of lethal cell injury in ischemic myocardium. Circulation 74, 1124-1136.

Muscat,J.E., Kleinman,W., Colosimo,S., Muir,A., Lazarus,P., Park,J., and Richie,J.P., Jr. (2004). Enhanced protein glutathiolation and oxidative stress in cigarette smokers. Free Radic. Biol. Med. 36, 464-470.

Mustapha,S., Kirshner,A., De,M.D., and Kirshenbaum,L.A. (2000). A direct requirement of nuclear factor-kappa B for suppression of apoptosis in ventricular myocytes. Am. J. Physiol Heart Circ. Physiol 279, H939-H945.

Nagy,N., Malik,G., Tosaki,A., Ho,Y.S., Maulik,N., and Das,D.K. (2008). Overexpression of glutaredoxin-2 reduces myocardial cell death by preventing both apoptosis and necrosis. J. Mol. Cell Cardiol. 44, 252-260.

Naumann,U., Wischhusen,J., Weit,S., Rieger,J., Wolburg,H., Massing,U., and Weller,M. (2004). Alkylphosphocholine-induced glioma cell death is BCL-X(L)-sensitive, caspase- independent and characterized by massive cytoplasmic vacuole formation. Cell Death. Differ. 11, 1326-1341.

Neimanis,S., Albig,W., Doenecke,D., and Kahle,J. (2007). Sequence elements in both subunits of the DNA fragmentation factor are essential for its nuclear transport. J. Biol. Chem. 282, 35821-35830.

Nikkola,M., Gleason,F.K., Saarinen,M., Joelson,T., Bjornberg,O., and Eklund,H. (1991). A putative glutathione-binding site in T4 glutaredoxin investigated by site-directed mutagenesis. J. Biol. Chem. 266, 16105-16112.

Nitahara,J.A., Cheng,W., Liu,Y., Li,B., Leri,A., Li,P., Mogul,D., Gambert,S.R., Kajstura,J., and Anversa,P. (1998). Intracellular calcium, DNase activity and myocyte apoptosis in aging Fischer 344 rats. J. Mol. Cell Cardiol. 30, 519-535.

Noguera,V., Walker,O., Rouhier,N., Jacquot,J.P., Krimm,I., and Lancelin,J.M. (2005). NMR reveals a novel glutaredoxin-glutaredoxin interaction interface. J. Mol. Biol. 353, 629-641.

302

Nonaka,K., Kume,N., Urata,Y., Seto,S., Kohno,T., Honda,S., Ikeda,S., Muroya,T., Ikeda,Y., Ihara,Y., Kita,T., and Kondo,T. (2007). Serum levels of S-glutathionylated proteins as a risk-marker for arteriosclerosis obliterans. Circ. J. 71, 100-105.

Nordstrand,K., slund,F., Holmgren,A., Otting,G., and Berndt,K.D. (1999). NMR structure of Escherichia coli glutaredoxin 3-glutathione mixed disulfide complex: implications for the enzymatic mechanism. J. Mol. Biol. 286, 541-552.

Nulton-Persson,A.C., Starke,D.W., Mieyal,J.J., and Szweda,L.I. (2003). Reversible inactivation of alpha-ketoglutarate dehydrogenase in response to alterations in the mitochondrial glutathione status. Biochemistry 42, 4235-4242.

Obin,M., Shang,F., Gong,X., Handelman,G., Blumberg,J., and Taylor,A. (1998). Redox regulation of ubiquitin-conjugating enzymes: mechanistic insights using the thiol-specific oxidant diamide. FASEB J. 12, 561-569.

Ogata,Y. and Takahashi,M. (2003). Bcl-xL as an antiapoptotic molecule for cardiomyocytes. Drug News Perspect. 16, 446-452.

Ojeda,L., Keller,G., Muhlenhoff,U., Rutherford,J.C., Lill,R., and Winge,D.R. (2006). Role of glutaredoxin-3 and glutaredoxin-4 in the iron regulation of the Aft1 transcriptional activator in Saccharomyces cerevisiae. J. Biol. Chem. 281, 17661-17669.

Okamoto,T., Akaike,T., Sawa,T., Miyamoto,Y., van der Vliet,A., and Maeda,H. (2001). Activation of matrix metalloproteinases by peroxynitrite-induced protein S- glutathiolation via disulfide S-oxide formation. J. Biol. Chem. 276, 29596-29602.

Okaya,T., Blanchard,J., Schuster,R., Kuboki,S., Husted,T., Caldwell,C.C., Zingarelli,B., Wong,H., Solomkin,J.S., and Lentsch,A.B. (2005). Age-dependent responses to hepatic ischemia/reperfusion injury. Shock 24, 421-427.

Okuda,M., Inoue,N., Azumi,H., Seno,T., Sumi,Y., Hirata,K., Kawashima,S., Hayashi,Y., Itoh,H., Yodoi,J., and Yokoyama,M. (2001). Expression of glutaredoxin in human coronary arteries: its potential role in antioxidant protection against atherosclerosis. Arterioscler. Thromb. Vasc. Biol. 21, 1483-1487.

Ono,M., Sawa,Y., Ryugo,M., Alechine,A.N., Shimizu,S., Sugioka,R., Tsujimoto,Y., Matsuda,H. (2005). BH4 peptide derivative from Bcl-xL attenuates ischemia/reperfusion injury thorough anti-apoptotic mechanism in rat hearts. Eur. J. Cardiothorac. Surg. 27, 117-121.

Ostrowski,M.C. and Kistler,W.S. (1980). Properties of a flavoprotein sulfhydryl oxidase from rat seminal vesicle secretion. Biochemistry 19, 2639-2645.

Pai,H.V., Starke,D.W., Lesnefsky,E.J., Hoppel,C.L., and Mieyal,J.J. (2007). What is the functional significance of the unique location of glutaredoxin 1 (GRx1) in the intermembrane space of mitochondria? Antioxid. Redox. Signal. 9, 2027-2033.

303

Pan,G., O'Rourke,K., and Dixit,V.M. (1998). Caspase-9, Bcl-XL, and Apaf-1 form a ternary complex. J. Biol. Chem. 273, 5841-5845.

Pan,S. and Berk,B.C. (2007). Glutathiolation regulates tumor necrosis factor-alpha- induced caspase-3 cleavage and apoptosis: key role for glutaredoxin in the death pathway. Circ. Res. 100, 213-219.

Papov,V.V., Gravina,S.A., Mieyal,J.J., and Biemann,K. (1994). The primary structure and properties of thioltransferase (glutaredoxin) from human red blood cells. Protein Sci. 3, 428-434.

Park,J.W. (1988). Reaction of S-nitrosoglutathione with sulfhydryl groups in protein. Biochem. Biophys. Res. Commun. 152, 916-920.

Park,J.W., Mieyal,J.J., Rhee,S.G., Chock,P.B. (2009). Deglutathionylation of 2-Cys peroxiredoxin is specifically catalyzed by sulfiredoxin. J. Biol. Chem. [Epub ahead of print, June 27]

Patel,H.H., Head,B.P., Petersen,H.N., Niesman,I.R., Huang,D., Gross,G.J., Insel,P.A., and Roth,D.M. (2006). Protection of adult rat cardiac myocytes from ischemic cell death: role of caveolar microdomains and delta-opioid receptors. Am. J. Physiol Heart Circ. Physiol 291, H344-H350.

Patel,J.R. and Brewer,G.J. (2008). Age-related differences in NFkappaB translocation and Bcl-2/Bax ratio caused by TNFalpha and Abeta42 promote survival in middle-age neurons and death in old neurons. Exp. Neurol. 213, 93-100.

Peltoniemi,M.J., Karala,A.R., Jurvansuu,J.K., Kinnula,V.L., and Ruddock,L.W. (2006a). Insights into deglutathionylation reactions. Different intermediates in the glutaredoxin and protein disulfide isomerase catalyzed reactions are defined by the gamma-linkage present in glutathione. J Biol. Chem. 281, 33107-33114.

Peltoniemi,M.J., Rytila,P.H., Harju,T.H., Soini,Y.M., Salmenkivi,K.M., Ruddock,L.W., and Kinnula,V.L. (2006b). Modulation of glutaredoxin in the lung and sputum of cigarette smokers and chronic obstructive pulmonary disease. Respir. Res. 7, 133.

Pfaff,E., Klingenberg,M., Ritt,E., and Vogell,W. (1968). [Correlation of the unspecific permeable mitochondrial space with the "intermembrane space"]. Eur. J. Biochem. 5, 222-232.

Phaneuf,S. and Leeuwenburgh,C. (2002). Cytochrome c release from mitochondria in the aging heart: a possible mechanism for apoptosis with age. Am. J. Physiol Regul. Integr. Comp Physiol 282, R423-R430.

Pimentel,D.R., Adachi,T., Ido,Y., Heibeck,T., Jiang,B., Lee,Y., Melendez,J.A., Cohen,R.A., and Colucci,W.S. (2006). Strain-stimulated hypertrophy in cardiac myocytes is mediated by reactive oxygen species-dependent Ras S-glutathiolation. J. Mol. Cell Cardiol. 41, 613-622.

304

Pineda-Molina,E., Klatt,P., Vazquez,J., Marina,A., Garcia,d.L., Perez-Sala,D., and Lamas,S. (2001). Glutathionylation of the p50 subunit of NF-kappaB: a mechanism for redox-induced inhibition of DNA binding. Biochemistry 40, 14134-14142.

Pintus,G., Tadolini,B., Posadino,A.M., Sanna,B., Debidda,M., Carru,C., Deiana,L., and Ventura,C. (2003). PKC/Raf/MEK/ERK signaling pathway modulates native-LDL- induced E2F-1 gene expression and endothelial cell proliferation. Cardiovasc. Res. 59, 934-944.

Poole,L.B., Karplus,P.A., and Claiborne,A. (2004). Protein sulfenic acids in redox signaling. Annu. Rev. Pharmacol. Toxicol. 44, 325-347.

Porcelli,A.M., Ghelli,A., Zanna,C., Pinton,P., Rizzuto,R., and Rugolo,M. (2005). pH difference across the outer mitochondrial membrane measured with a green fluorescent protein mutant. Biochem. Biophys. Res. Commun. 326, 799-804.

Qanungo,S., Starke,D.W., Pai,H.V., Mieyal,J.J., and Nieminen,A.L. (2007). Glutathione supplementation potentiates hypoxic apoptosis by S-glutathionylation of p65-NFkappaB. J. Biol. Chem. 282, 18427-18436.

Qiao,F., Xing,K., Liu,A., Ehlers,N., Raghavachari,N., and Lou,M.F. (2001). Human lens thioltransferase: cloning, purification, and function. Invest Ophthalmol. Vis. Sci. 42, 743- 751.

Quinlan,P.T., Thomas,A.P., Armston,A.E., Halestrap,A.P. (1983). Measurement of the intramitochondrial volume in hepatocytes without cell disruption and its elevation by hormones and valinomycin. Biochem. J. 214, 395-404.

Raghavachari,N., Qiao,F., and Lou,M.F. (1999). Does glutathione-S-transferase dethiolate lens protein-thiol mixed disulfides?-A comparative study with thioltransferase. Exp. Eye Res. 68, 715-724.

Ralat,L.A., Manevich,Y., Fisher,A.B., and Colman,R.F. (2006). Direct evidence for the formation of a complex between 1-cysteine peroxiredoxin and glutathione S-transferase pi with activity changes in both enzymes. Biochemistry 45, 360-372.

Reynaert,N.L., van der Vliet,A., Guala,A.S., McGovern,T., Hristova,M., Pantano,C., Heintz,N.H., Heim,J., Ho,Y.S., Matthews,D.E., Wouters,E.F., and Janssen- Heininger,Y.M. (2006). Dynamic redox control of NF-kappaB through glutaredoxin- regulated S-glutathionylation of inhibitory kappaB kinase beta. Proc. Natl. Acad. Sci. U. S. A 103, 13086-13091.

Rhee,S.G., Chae,H.Z., and Kim,K. (2005). Peroxiredoxins: a historical overview and speculative preview of novel mechanisms and emerging concepts in cell signaling. Free Radic. Biol. Med. 38, 1543-1552.

305

Rich,M.W., Bosner,M.S., Chung,M.K., Shen,J., and McKenzie,J.P. (1992). Is age an independent predictor of early and late mortality in patients with acute myocardial infarction? Am. J. Med. 92, 7-13.

Rikans,L.E. and Hornbrook,K.R. (1998). Thiol-disulfide exchange systems in the liver of aging Fischer 344 rats. Gerontology 44, 72-77.

Rodriguez-Manzaneque,M.T., Ros,J., Cabiscol,E., Sorribas,A., and Herrero,E. (1999). Grx5 glutaredoxin plays a central role in protection against protein oxidative damage in Saccharomyces cerevisiae. Mol. Cell Biol. 19, 8180-8190.

Rodriguez-Manzaneque,M.T., Tamarit,J., Belli,G., Ros,J., and Herrero,E. (2002). Grx5 is a mitochondrial glutaredoxin required for the activity of iron/sulfur enzymes. Mol. Biol. Cell 13, 1109-1121.

Rong,Y. and Distelhorst,C.W. (2008). Bcl-2 protein family members: versatile regulators of calcium signaling in cell survival and apoptosis. Annu. Rev. Physiol 70, 73-91.

Rong,Y.P., Aromolaran,A.S., Bultynck,G., Zhong,F., Li,X., McColl,K., Matsuyama,S., Herlitze,S., Roderick,H.L., Bootman,M.D., Mignery,G.A., Parys,J.B., De,S.H., and Distelhorst,C.W. (2008). Targeting Bcl-2-IP3 receptor interaction to reverse Bcl-2's inhibition of apoptotic calcium signals. Mol. Cell 31, 255-265.

Rossi,R., Giustarini,D., Dalle-Donne,I., and Milzani,A. (2006). Protein S- glutathionylation and platelet anti-aggregating activity of disulfiram. Biochem. Pharmacol. 72, 608-615.

Rouhier,N., Gelhaye,E., and Jacquot,J.P. (2002a). Exploring the active site of plant glutaredoxin by site-directed mutagenesis. FEBS Lett. 511, 145-149.

Rouhier,N., Gelhaye,E., and Jacquot,J.P. (2002b). Glutaredoxin-dependent peroxiredoxin from poplar: protein-protein interaction and catalytic mechanism. J. Biol. Chem. 277, 13609-13614.

Rouhier,N., Gelhaye,E., and Jacquot,J.P. (2004). Plant glutaredoxins: still mysterious reducing systems. Cell Mol. Life Sci. 61, 1266-1277.

Rouhier,N., Gelhaye,E., Sautiere,P.E., Brun,A., Laurent,P., Tagu,D., Gerard,J., De,F.E., Meyer,Y., and Jacquot,J.P. (2001). Isolation and characterization of a new peroxiredoxin from poplar sieve tubes that uses either glutaredoxin or thioredoxin as a proton donor. Plant Physiol 127, 1299-1309.

Rouhier,N., Unno,H., Bandyopadhyay,S., Masip,L., Kim,S.K., Hirasawa,M., Gualberto,J.M., Lattard,V., Kusunoki,M., Knaff,D.B., Georgiou,G., Hase,T., Johnson,M.K., and Jacquot,J.P. (2007). Functional, structural, and spectroscopic characterization of a glutathione-ligated [2Fe-2S] cluster in poplar glutaredoxin C1. Proc. Natl. Acad. Sci. U. S. A 104, 7379-7384.

306

Saaranen,M., Salo,K., Latva-Ranta,M., Kinnula,V., and Ruddock,L.W. (2009). THE C- TERMINAL ACTIVE SITE CYSTEINE OF ESCHERICHIA COLI GLUTAREDOXIN 1 DETERMINES THE GLUTATHIONE SPECIFICITY OF THE SECOND STEP OF PEPTIDE DEGLUTATHIONYLATION. Antioxid. Redox. Signal.

Saavedra,J.E., Billiar,T.R., Williams,D.L., Kim,Y.M., Watkins,S.C., and Keefer,L.K. (1997). Targeting nitric oxide (NO) delivery in vivo. Design of a liver-selective NO donor prodrug that blocks tumor necrosis factor-alpha-induced apoptosis and toxicity in the liver. J. Med. Chem. 40, 1947-1954.

Sadidi,M., Geddes,T.J., and Kuhn,D.M. (2005). S-thiolation of tyrosine hydroxylase by reactive nitrogen species in the presence of cysteine or glutathione. Antioxid. Redox. Signal. 7, 863-869.

Sagemark,J., Elgan,T.H., Burglin,T.R., Johansson,C., Holmgren,A., and Berndt,K.D. (2007). Redox properties and evolution of human glutaredoxins. Proteins 68, 879-892.

Sakamoto,K.M. and Frank,D.A. (2009). CREB in the pathophysiology of cancer: implications for targeting transcription factors for cancer therapy. Clin. Cancer Res. 15, 2583-2587.

Salmeen,A., Andersen,J.N., Myers,M.P., Meng,T.C., Hinks,J.A., Tonks,N.K., and Barford,D. (2003). Redox regulation of protein tyrosine phosphatase 1B involves a sulphenyl-amide intermediate. Nature 423, 769-773.

Samudio,I., Konopleva,M., Hail,N., Jr., Shi,Y.X., McQueen,T., Hsu,T., Evans,R., Honda,T., Gribble,G.W., Sporn,M., Gilbert,H.F., Safe,S., and Andreeff,M. (2005). 2- Cyano-3,12-dioxooleana-1,9-dien-28-imidazolide (CDDO-Im) directly targets mitochondrial glutathione to induce apoptosis in pancreatic cancer. J. Biol. Chem. 280, 36273-36282.

Sanada,S., Asanuma,H., Tsukamoto,O., Minamino,T., Node,K., Takashima,S., Fukushima,T., Ogai,A., Shinozaki,Y., Fujita,M., Hirata,A., Okuda,H., Shimokawa,H., Tomoike,H., Hori,M., and Kitakaze,M. (2004). Protein kinase A as another mediator of ischemic preconditioning independent of protein kinase C. Circulation 110, 51-57.

Sanchez,G., Escobar,M., Pedrozo,Z., Macho,P., Domenech,R., Hartel,S., Hidalgo,C., and Donoso,P. (2008). Exercise and tachycardia increase NADPH oxidase and ryanodine receptor-2 activity: possible role in cardioprotection. Cardiovasc. Res. 77, 380-386.

Sanchez,G., Pedrozo,Z., Domenech,R.J., Hidalgo,C., and Donoso,P. (2005). Tachycardia increases NADPH oxidase activity and RyR2 S-glutathionylation in ventricular muscle. J. Mol. Cell Cardiol. 39, 982-991.

Santa,M.C. and Machado,A. (1986). Age and sex related differences in some rat renal NADPH-consuming detoxification enzymes. Arch. Gerontol. Geriatr. 5, 235-247.

Schoneich,C. (1995). Kinetics of thiol reactions. Methods Enzymol. 251, 45-55.

307

Sevilla,L., Zaldumbide,A., Pognonec,P., and Boulukos,K.E. (2001). Transcriptional regulation of the bcl-x gene encoding the anti-apoptotic Bcl-xL protein by Ets, Rel/NFkappaB, STAT and AP1 transcription factor families. Histol. Histopathol. 16, 595-601.

Sha,S., Minakuchi,K., Higaki,N., Sato,K., Ohtsuki,K., Kurata,A., Yoshikawa,H., Kotaru,M., Masumura,T., Ichihara,K., and Tanaka,K. (1997). Purification and characterization of glutaredoxin (thioltransferase) from rice (Oryza sativa L.). J. Biochem. 121, 842-848.

Shelton,M.D., Chock,P.B., and Mieyal,J.J. (2005). Glutaredoxin: role in reversible protein s-glutathionylation and regulation of redox signal transduction and protein translocation. Antioxid. Redox. Signal. 7, 348-366.

Shelton,M.D., Distler,A.M., Kern,T.S., and Mieyal,J.J. (2009). Glutaredoxin regulates autocrine and paracrine proinflammatory responses in retinal glial (muller) cells. J. Biol. Chem. 284, 4760-4766.

Shelton,M.D., Kern,T.S., and Mieyal,J.J. (2007). Glutaredoxin regulates nuclear factor kappa-B and intercellular adhesion molecule in Muller cells: model of diabetic retinopathy. J Biol. Chem. 282, 12467-12474.

Shelton,M.D. and Mieyal,J.J. (2008). Regulation by reversible S-glutathionylation: molecular targets implicated in inflammatory diseases. Mol. Cells 25, 332-346.

Shenton,D., Perrone,G., Quinn,K.A., Dawes,I.W., and Grant,C.M. (2002). Regulation of protein S-thiolation by glutaredoxin 5 in the yeast Saccharomyces cerevisiae. J. Biol. Chem. 277, 16853-16859.

Shi,J., Vlamis-Gardikas,A., Aslund,F., Holmgren,A., and Rosen,B.P. (1999). Reactivity of glutaredoxins 1, 2, and 3 from Escherichia coli shows that glutaredoxin 2 is the primary hydrogen donor to ArsC-catalyzed arsenate reduction. J. Biol. Chem. 274, 36039-36042.

Shimizu,S., Eguchi,Y., Kamiike,W., Funahashi,Y., Mignon,A., Lacronique,V., Matsuda,H., and Tsujimoto,Y. (1998). Bcl-2 prevents apoptotic mitochondrial dysfunction by regulating proton flux. Proc. Natl. Acad. Sci. U. S. A 95, 1455-1459.

Shoneich, C. Thiyl Radicals, Perththiyl radicals, and Oxidative Reactions. Packer, L and Cadenas, E. 21-47. 1995. Marcel Dekker, Inc. Biothiols in Health and Disease. Ref Type: Serial (Book,Monograph)

Silva,G.M., Netto,L.E., Discola,K.F., Piassa-Filho,G.M., Pimenta,D.C., Barcena,J.A., and Demasi,M. (2008). Role of glutaredoxin 2 and cytosolic thioredoxins in cysteinyl-based redox modification of the 20S proteasome. FEBS J. 275, 2942-2955.

308

Singh,S.P., Wishnok,J.S., Keshive,M., Deen,W.M., and Tannenbaum,S.R. (1996). The chemistry of the S-nitrosoglutathione/glutathione system. Proc. Natl. Acad. Sci. U. S. A 93, 14428-14433.

Sohal,R.S., Farmer,K.J., and Allen,R.G. (1987). Correlates of longevity in two strains of the housefly, Musca domestica. Mech. Ageing Dev. 40, 171-179.

Song,J.J., Rhee,J.G., Suntharalingam,M., Walsh,S.A., Spitz,D.R., and Lee,Y.J. (2002). Role of glutaredoxin in metabolic oxidative stress. Glutaredoxin as a sensor of oxidative stress mediated by H2O2. J. Biol. Chem. 277, 46566-46575.

Srinivasan,U., Mieyal,P.A., and Mieyal,J.J. (1997). pH profiles indicative of rate-limiting nucleophilic displacement in thioltransferase catalysis. Biochemistry 36, 3199-3206.

Starke,D.W., Chen,Y., Bapna,C.P., Lesnefsky,E.J., and Mieyal,J.J. (1997). Sensitivity of protein sulfhydryl repair enzymes to oxidative stress. Free Radic. Biol. Med. 23, 373-384.

Starke,D.W., Chock,P.B., and Mieyal,J.J. (2003). Glutathione-thiyl radical scavenging and transferase properties of human glutaredoxin (thioltransferase). Potential role in redox signal transduction. J. Biol. Chem. 278, 14607-14613.

Stavreus-Evers,A., Masironi,B., Landgren,B.M., Holmgren,A., Eriksson,H., and Sahlin,L. (2002). Immunohistochemical localization of glutaredoxin and thioredoxin in human endometrium: a possible association with pinopodes. Mol. Hum. Reprod. 8, 546- 551.

Sturm,B., Twaroch,T., Knapitsch,B., Czingraber,S., Ternes,N., Goldenberg,H., and Scheiber-Mojdehkar,B. (2006). Differential response of iron metabolism to oxidative stress generated by antimycin A and nitrofurantoin. Biochimie 88, 575-581.

Su,D. and Gladyshev,V.N. (2004). Alternative splicing involving the thioredoxin reductase module in mammals: a glutaredoxin-containing thioredoxin reductase 1. Biochemistry 43, 12177-12188.

Su,D., Novoselov,S.V., Sun,Q.A., Moustafa,M.E., Zhou,Y., Oko,R., Hatfield,D.L., and Gladyshev,V.N. (2005). Mammalian selenoprotein thioredoxin-glutathione reductase. Roles in disulfide bond formation and sperm maturation. J. Biol. Chem. 280, 26491- 26498.

Suh,J.H., Heath,S.H., and Hagen,T.M. (2003). Two subpopulations of mitochondria in the aging rat heart display heterogenous levels of oxidative stress. Free Radic. Biol. Med. 35, 1064-1072.

Sullivan,D.M., Wehr,N.B., Fergusson,M.M., Levine,R.L., and Finkel,T. (2000). Identification of oxidant-sensitive proteins: TNF-alpha induces protein glutathiolation. Biochemistry 39, 11121-11128.

309

Sun,C., Berardi,M.J., and Bushweller,J.H. (1998). The NMR solution structure of human glutaredoxin in the fully reduced form. J. Mol. Biol. 280, 687-701.

Sun,Q.A., Kirnarsky,L., Sherman,S., and Gladyshev,V.N. (2001). Selenoprotein oxidoreductase with specificity for thioredoxin and glutathione systems. Proc. Natl. Acad. Sci. U. S. A 98, 3673-3678.

Sun,Q.A., Su,D., Novoselov,S.V., Carlson,B.A., Hatfield,D.L., and Gladyshev,V.N. (2005). Reaction mechanism and regulation of mammalian thioredoxin/glutathione reductase. Biochemistry 44, 14528-14537.

Sundaram,S., Rathinasabapathi,B., Ma,L.Q., and Rosen,B.P. (2008). An arsenate- activated glutaredoxin from the arsenic hyperaccumulator fern Pteris vittata L. regulates intracellular arsenite. J. Biol. Chem. 283, 6095-6101.

Swift,L.L. (1996). Role of the Golgi apparatus in the phosphorylation of apolipoprotein B. J Biol. Chem. 271, 31491-31495.

Sykes,M.C., Mowbray,A.L., and Jo,H. (2007). Reversible glutathiolation of caspase-3 by glutaredoxin as a novel redox signaling mechanism in tumor necrosis factor-alpha- induced cell death. Circ. Res. 100, 152-154.

Szajewski,R.P. and Whitesides,G.M. (1980). Rate Constants and Equilibrium-Constants for Thiol-Disulfide Interchange Reactions Involving Oxidized Glutathione. Journal of the American Chemical Society 102, 2011-2026.

Tamarit,J., Belli,G., Cabiscol,E., Herrero,E., and Ros,J. (2003). Biochemical characterization of yeast mitochondrial Grx5 monothiol glutaredoxin. J. Biol. Chem. 278, 25745-25751.

Tanaka,T., Nakamura,H., Nishiyama,A., Hosoi,F., Masutani,H., Wada,H., and Yodoi,J. (2000). Redox regulation by thioredoxin superfamily; protection against oxidative stress and aging. Free Radic. Res. 33, 851-855.

Tani,M., Suganuma,Y., Hasegawa,H., Shinmura,K., Ebihara,Y., Hayashi,Y., Guo,X., and Takayama,M. (1997). Decrease in ischemic tolerance with aging in isolated perfused Fischer 344 rat hearts: relation to increases in intracellular Na+ after ischemia. J. Mol. Cell Cardiol. 29, 3081-3089.

Tantini,B., Fiumana,E., Cetrullo,S., Pignatti,C., Bonavita,F., Shantz,L.M., Giordano,E., Muscari,C., Flamigni,F., Guarnieri,C., Stefanelli,C., and Caldarera,C.M. (2006). Involvement of polyamines in apoptosis of cardiac myoblasts in a model of simulated ischemia. J. Mol. Cell Cardiol. 40, 775-782.

Taylor,E.R., Hurrell,F., Shannon,R.J., Lin,T.K., Hirst,J., and Murphy,M.P. (2003). Reversible glutathionylation of complex I increases mitochondrial superoxide formation. J. Biol. Chem. 278, 19603-19610.

310

Tew,K.D. (2007). Redox in redux: Emergent roles for glutathione S-transferase P (GSTP) in regulation of cell signaling and S-glutathionylation. Biochem. Pharmacol. 73, 1257- 1269.

Thomas,J.A., Poland,B., and Honzatko,R. (1995). Protein sulfhydryls and their role in the antioxidant function of protein S-thiolation. Arch. Biochem. Biophys. 319, 1-9.

Thorpe,C., Hoober,K.L., Raje,S., Glynn,N.M., Burnside,J., Turi,G.K., and Coppock,D.L. (2002). Sulfhydryl oxidases: emerging catalysts of protein disulfide bond formation in eukaryotes. Arch. Biochem. Biophys. 405, 1-12.

Tomasini,R., Tsuchihara,K., Wilhelm,M., Fujitani,M., Rufini,A., Cheung,C.C., Khan,F., Itie-Youten,A., Wakeham,A., Tsao,M.S., Iovanna,J.L., Squire,J., Jurisica,I., Kaplan,D., Melino,G., Jurisicova,A., and Mak,T.W. (2008). TAp73 knockout shows genomic instability with infertility and tumor suppressor functions. Genes Dev. 22, 2677-2691.

Townsend,D.M., Findlay,V.J., Fazilev,F., Ogle,M., Fraser,J., Saavedra,J.E., Ji,X., Keefer,L.K., and Tew,K.D. (2006). A glutathione S-transferase pi-activated prodrug causes kinase activation concurrent with S-glutathionylation of proteins. Mol. Pharmacol. 69, 501-508.

Ungvari,Z., Orosz,Z., Labinskyy,N., Rivera,A., Xiangmin,Z., Smith,K., and Csiszar,A. (2007). Increased mitochondrial H2O2 production promotes endothelial NF-kappaB activation in aged rat arteries. Am. J. Physiol Heart Circ. Physiol 293, H37-H47.

Urata,Y., Ihara,Y., Murata,H., Goto,S., Koji,T., Yodoi,J., Inoue,S., and Kondo,T. (2006). 17Beta-estradiol protects against oxidative stress-induced cell death through the glutathione/glutaredoxin-dependent redox regulation of Akt in myocardiac H9c2 cells. J. Biol. Chem. 281, 13092-13102. van Montfort,R.L., Congreve,M., Tisi,D., Carr,R., and Jhoti,H. (2003). Oxidation state of the active-site cysteine in protein tyrosine phosphatase 1B. Nature 423, 773-777.

Vander Heiden,M.G., Chandel,N.S., Schumacker,P.T., and Thompson,C.B. (1999). Bcl- xL prevents cell death following growth factor withdrawal by facilitating mitochondrial ATP/ADP exchange. Mol. Cell 3, 159-167.

Venters,H.D., Jr., Bonilla,L.E., Jensen,T., Garner,H.P., Bordayo,E.Z., Najarian,M.M., Ala,T.A., Mason,R.P., and Frey,W.H. (1997). Heme from Alzheimer's brain inhibits muscarinic receptor binding via thiyl radical generation. Brain Res. 764, 93-100.

Vlamis-Gardikas,A., Aslund,F., Spyrou,G., Bergman,T., and Holmgren,A. (1997). Cloning, overexpression, and characterization of glutaredoxin 2, an atypical glutaredoxin from Escherichia coli. J. Biol. Chem. 272, 11236-11243.

Vlamis-Gardikas,A., Potamitou,A., Zarivach,R., Hochman,A., and Holmgren,A. (2002). Characterization of Escherichia coli null mutants for glutaredoxin 2. J. Biol. Chem. 277, 10861-10868.

311

Voehringer,D.W. and Meyn,R.E. (2000). Redox aspects of Bcl-2 function. Antioxid. Redox. Signal. 2, 537-550.

Wang,H.D., Xu,S., Johns,D.G., Du,Y., Quinn,M.T., Cayatte,A.J., and Cohen,R.A. (2001a). Role of NADPH oxidase in the vascular hypertrophic and oxidative stress response to angiotensin II in mice. Circ. Res. 88, 947-953.

Wang,J., Boja,E.S., Tan,W., Tekle,E., Fales,H.M., English,S., Mieyal,J.J., and Chock,P.B. (2001b). Reversible glutathionylation regulates actin polymerization in A431 cells. J. Biol. Chem. 276, 47763-47766.

Wang,J., Pan,S., and Berk,B.C. (2007). Glutaredoxin mediates Akt and eNOS activation by flow in a glutathione reductase-dependent manner. Arterioscler. Thromb. Vasc. Biol. 27, 1283-1288.

Wang,J., Tekle,E., Oubrahim,H., Mieyal,J.J., Stadtman,E.R., and Chock,P.B. (2003). Stable and controllable RNA interference: Investigating the physiological function of glutathionylated actin. Proc. Natl. Acad. Sci. U. S. A 100, 5103-5106.

Wang,L., Guo,Y., Huang,W.J., Ke,X., Poyet,J.L., Manji,G.A., Merriam,S., Glucksmann,M.A., DiStefano,P.S., Alnemri,E.S., and Bertin,J. (2001c). Card10 is a novel caspase recruitment domain/membrane-associated guanylate kinase family member that interacts with BCL10 and activates NF-kappa B. J. Biol. Chem. 276, 21405-21409.

Wang,X., Zhang,J., Kim,H.P., Wang,Y., Choi,A.M., and Ryter,S.W. (2004). Bcl-XL disrupts death-inducing signal complex formation in plasma membrane induced by hypoxia/reoxygenation. FASEB J. 18, 1826-1833.

Wang,Y., Qiao,M., Mieyal,J.J., Asmis,L.M., and Asmis,R. (2006). Molecular mechanism of glutathione-mediated protection from oxidized low-density lipoprotein-induced cell injury in human macrophages: role of glutathione reductase and glutaredoxin. Free Radic. Biol. Med. 41, 775-785.

Wardman, P. Reactions of Thiyl Radicals. Packer, L and Cadenas, E. 1-19. 1995. Marcel Dekker, Inc. Ref Type: Serial (Book,Monograph)

Wardman,P. and von Sonntag,C. (1995). Kinetic factors that control the fate of thiyl radicals in cells. Methods Enzymol. 251, 31-45.

Webster,K.A., Graham,R.M., Thompson,J.W., Spiga,M.G., Frazier,D.P., Wilson,A., and Bishopric,N.H. (2006). Redox stress and the contributions of BH3-only proteins to infarction. Antioxid. Redox. Signal. 8, 1667-1676.

Weisleder,N., Taffet,G.E., and Capetanaki,Y. (2004). Bcl-2 overexpression corrects mitochondrial defects and ameliorates inherited desmin null cardiomyopathy. Proc. Natl. Acad. Sci. U. S. A 101, 769-774.

312

Wells,W.W., Rocque,P.A., Xu,D.P., Meyer,E.B., Charamella,L.J., and Dimitrov,N.V. (1995). Ascorbic acid and cell survival of adriamycin resistant and sensitive MCF-7 breast tumor cells. Free Radic. Biol. Med. 18, 699-708.

Willems,L., Zatta,A., Holmgren,K., Ashton,K.J., and Headrick,J.P. (2005). Age-related changes in ischemic tolerance in male and female mouse hearts. J. Mol. Cell Cardiol. 38, 245-256.

Wingert,R.A., Galloway,J.L., Barut,B., Foott,H., Fraenkel,P., Axe,J.L., Weber,G.J., Dooley,K., Davidson,A.J., Schmid,B., Paw,B.H., Shaw,G.C., Kingsley,P., Palis,J., Schubert,H., Chen,O., Kaplan,J., and Zon,L.I. (2005). Deficiency of glutaredoxin 5 reveals Fe-S clusters are required for vertebrate haem synthesis. Nature 436, 1035-1039.

Witte,S., Villalba,M., Bi,K., Liu,Y., Isakov,N., and Altman,A. (2000). Inhibition of the c- Jun N-terminal kinase/AP-1 and NF-kappaB pathways by PICOT, a novel protein kinase C-interacting protein with a thioredoxin homology domain. J. Biol. Chem. 275, 1902- 1909.

Yang,Y., Jao,S., Nanduri,S., Starke,D.W., Mieyal,J.J., and Qin,J. (1998). Reactivity of the human thioltransferase (glutaredoxin) C7S, C25S, C78S, C82S mutant and NMR solution structure of its glutathionyl mixed disulfide intermediate reflect catalytic specificity. Biochemistry 37, 17145-17156.

Yang,Y.F. and Wells,W.W. (1991). Catalytic mechanism of thioltransferase. J. Biol. Chem. 266, 12766-12771.

Yoshitake,S., Nanri,H., Fernando,M.R., and Minakami,S. (1994). Possible differences in the regenerative roles played by thioltransferase and thioredoxin for oxidatively damaged proteins. J. Biochem. (Tokyo) 116, 42-46.

Yu,J., Zhang,N.N., Yin,P.D., Cui,P.X., and Zhou,C.Z. (2008). Glutathionylation- triggered conformational changes of glutaredoxin Grx1 from the yeast Saccharomyces cerevisiae. Proteins 72, 1077-1083.

Zhang,H., Tao,L., Jiao,X., Gao,E., Lopez,B.L., Christopher,T.A., Koch,W., and Ma,X.L. (2007). Nitrative thioredoxin inactivation as a cause of enhanced myocardial ischemia/reperfusion injury in the aging heart. Free Radic. Biol. Med. 43, 39-47.

Zhao,T.C. and Kukreja,R.C. (2002). Late preconditioning elicited by activation of adenosine A(3) receptor in heart: role of NF- kappa B, iNOS and mitochondrial K(ATP) channel. J. Mol. Cell Cardiol. 34, 263-277.

Zidar,N., Jera,J., Maja,J., and Dusan,S. (2007). Caspases in myocardial infarction. Adv. Clin. Chem. 44, 1-33.

Ziegler,D.M. (1985). Role of reversible oxidation-reduction of enzyme thiols-disulfides in metabolic regulation. Annu. Rev. Biochem. 54, 305-329.

313

Zimmermann,A.K., Loucks,F.A., Schroeder,E.K., Bouchard,R.J., Tyler,K.L., and Linseman,D.A. (2007). Glutathione binding to the Bcl-2 homology-3 domain groove: a molecular basis for Bcl-2 antioxidant function at mitochondria. J. Biol. Chem. 282, 29296-29304.

314